Arabidopsis FAB1/PIKfyve Proteins Are Essential for
Development of Viable Pollen[W]
Paul Whitley1, Steven Hinz, and James Doughty1*
Department of Biology and Biochemistry, University of Bath, Bath, Somerset BA2 7AY, United Kingdom
Phosphatidylinositol 3,5-bisphosphate [PtdIns(3,5)P2] is a phospholipid that has a role in controlling membrane trafficking
events in yeast and animal cells. The function of this lipid in plants is unknown, although its synthesis has been shown to be
up-regulated upon osmotic stress in plant cells. PtdIns(3,5)P2 is synthesized by the PIKfyve/Fab1 family of proteins, with two
orthologs, FAB1A and FAB1B, being present in Arabidopsis (Arabidopsis thaliana). In this study, we attempt to address the role
of this lipid by analyzing the phenotypes of plants mutated in FAB1A and FAB1B. It was not possible to generate plants
homozygous for mutations in both genes, although single mutants were isolated. Both homozygous single mutant plant lines
exhibited a leaf curl phenotype that was more marked in FAB1B mutants. Genetic transmission analysis revealed that failure to
generate double mutant lines was entirely due to inviability of pollen carrying mutant alleles of both FAB1A and FAB1B. This
pollen displayed severe defects in vacuolar reorganization following the first mitotic division of development. The presence of
abnormally large vacuoles in pollen at the tricellular stage resulted in the collapse of the majority of grains carrying both
mutant alleles. This demonstrates a crucial role for PtdIns(3,5)P2 in modulating the dynamics of vacuolar rearrangement
essential for successful pollen development. Taken together, our results are consistent with PtdIns(3,5)P2 production being
central to cellular responses to changes in osmotic conditions.
Phosphoinositides make up a minor fraction of total
membrane lipids in all eukaryotic organisms. Their
production is spatially restricted to the cytoplasmic
leaflet of specific organellar membranes and temporally regulated by phosphatidylinositol (PtdIns) kinases and phosphatases. Three of the five hydoxyl
groups of PtdIns can be phosphorylated, either singly
or combinatorially, to produce seven different phosphoinositides. These different phosphoinositides
can recruit and/or activate proteins with specific
phosphoinositide-binding domains and have been
implicated in the regulation of many important cellular functions, including membrane trafficking, cell
growth, and cytoskeleton remodeling (Di Paolo and
De Camilli, 2006).
In animal cells, phosphorylation at the 3 position of
PtdIns and its phosphorylated derivatives can be
carried out by three different classes of PtdIns 3-kinase
(classes I–III; Cantley, 2002). Plants and yeast only
have class III PtdIns 3-kinases that are orthologs of
the Saccharomyces cerevisiae protein Vps34p (MuellerRoeber and Pical, 2002). Vps34p orthologs are thought
to use PtdIns as their sole lipid substrate and produce
PtdIns 3-phosphate (PtdIns3P). PtdIns3P is involved
in endosomal/lysosomal protein sorting in eukaryotic
1
These authors contributed equally to the article.
* Corresponding author; e-mail [email protected].
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy
described in the Instructions for Authors (www.plantphysiol.org) is:
James Doughty ([email protected]).
[W]
The online version of this article contains Web-only data.
www.plantphysiol.org/cgi/doi/10.1104/pp.109.146159
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cells in addition to cellular signaling events (Backer,
2008).
In plants, PtdIns3P is essential for normal growth
and development. Arabidopsis (Arabidopsis thaliana)
plants carrying a VPS34 antisense construct have
severe developmental defects (Welters et al., 1994).
Furthermore, using pharmacological inhibitors of
PtdIns3P production and analysis of transgenic plants
defective in downstream signaling from PtdIns3P, it
has been shown that this lipid has a role to play in
many diverse physiological processes, such as root
hair growth (Lee et al., 2008a). The phenotypes observed in studies of PtdIns3P function in plants are
consistent with a role in endosomal and vacuolar
trafficking in plants (Kim et al., 2001; Lee et al.,
2008a), as in other eukaryotes. Recently, an attempt
to generate vps34 homozygous mutant plant lines was
unsuccessful due to failure of the mutant vps34 allele
to transmit through the male germ line (Lee et al.,
2008b).
Importantly, PtdIns3P is the precursor to another phosphoinositide, PtdIns 3,5-bisphosphate [PtdIns(3,5)P2],
which also has vital roles in endosomal trafficking in
eukaryotes (Dove et al., 2009). Thus, it is possible that
some of the effects in plants attributed to PtdIns3P in
previous studies may actually be due to an inability of
cells to produce PtdIns(3,5)P2. PtdIns(3,5)P2 is produced by the PtdIns3P 5-kinases PIKfyve and Fab1p in
animal and yeast cells, respectively. PIKfyve/Fab1p
proteins have an N-terminal FYVE domain necessary
for binding to PtdIns3P-containing membranes, a central Cpn60_TCP1 (for HSP chaperonin T complex 1)
homology domain, and a C-terminal kinase domain.
In Arabidopsis, there are a number of genes encoding
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Role of PtdIns(3)P 5-Kinase in Pollen Development
putative Fab1p homologs, but only two of them,
FAB1A (At4g33240) and FAB1B (At3g14270), encode
proteins having FYVE domains at their N termini
(Mueller-Roeber and Pical, 2002). It is likely that these
proteins are PtdIns3P 5-kinases in Arabidopsis. Despite the importance of PtdIns(3,5)P2 in yeast and
animals, very little is known about its function in
plants. However, it has been shown that hyperosmotic
stress can induce the rapid synthesis of PtdIns(3,5)P2
in cell suspension cultures from a number of plant
species (Meijer and Munnik, 2003) and in pollen tubes
from tobacco (Nicotiana tabacum; Zonia and Munnik,
2004). This production is consistent with a requirement for PtdIns(3,5)P2 in vacuolar membrane reorganization, as water moves from the vacuole to the
cytosol upon cells being placed under hyperosmotic
stress. In animal cells, defective PtdIns(3,5)P2 production leads to cytoplasmic vacuolation of endosomederived membranes (Ikonomov et al., 2001; Jefferies
et al., 2008). It seems that there is a general requirement in all eukaryotes for PtdIns(3,5)P2 production in
endomembrane remodeling. This remodeling could be
mediated by proteins that bind to PtdIns(3,5)P2. A
number of candidates have been identified, including
yeast Svp1p (Dove et al., 2004), its mammalian homolog WIP149 (Jeffries et al., 2004), CHMP3 (Whitley
et al., 2003), and Ent3p (Friant et al., 2003).
In this study, we aimed to further investigate the
role of PtdIns(3,5)P2 in plant physiology and the
function of PIKfyve/Fab1p orthologs in Arabidopsis
by generating mutant plant lines homozygous for
T-DNA insertions in both FAB1A and FAB1B. We failed
to generate double homozygous fab1a/fab1b knockout
plants but observed subtle phenotypes in both fab1a
and fab1b single homozygous plants. The data show
that pollen with a fab1a/fab1b genotype has an abnormal vacuolar phenotype and does not contribute to the
next generation. Our data are consistent with the
hypothesis that the male gametophytic defect observed in vps34 mutant pollen (Lee et al., 2008b) is
due to an inability of this pollen to generate PtdIns(3,5)P2
and is not a direct result of the lack of PtdIns3P.
RESULTS
Phylogeny and Expression of PtdIns3P 5-Kinases
in Arabidopsis
The availability of the complete Arabidopsis genome sequence has allowed the identification of
enzymes involved in phosphoinositide metabolism
based on comparison with the wealth of data from
studies in animals and yeast (for review, see MuellerRoeber and Pical, 2002). It has been proposed that
there are four proteins in Arabidopsis that are putative
homologs of PIKfyve and Fab1p, the enzymes responsible for the conversion of PtdIns3P to PtdIns(3,5)P2
(Fig. 1A), in animals and yeast, respectively (MuellerRoeber and Pical, 2002). Animals and yeast contain a
single copy of genes encoding PIKfyve/Fab1p.
The genes encoding the four putative PIKfyve/
Fab1p homologs in Arabidopsis are At4g33240
(FAB1A), At3g14270 (FAB1B), At1g71010 (FAB1C),
and At1g34260 (FAB1D). However, only FAB1A and
FAB1B encode proteins that contain predicted FYVE
domains at their N termini and therefore have the
same domain organization as PIKfyve/Fab1p (Fig.
1B). Phylogenetic analysis indicates that proteins similar to FAB1C and FAB1D, without a FYVE domain, are
likely to exist in all higher plant species and that these
proteins cluster in a plant-specific group distinct from
plant FYVE domain-containing proteins (Fig. 1C). We
propose that FAB1A and FAB1B encode bona fide
PtdIns3P 5-kinases, whereas the role of FAB1C and
FAB1D in the production of PtdIns(3,5)P2 is more
debatable. FAB1C and FAB1D are predicted to encode
polypeptides with PtdIns3P 5-kinase activity, but as
they lack a FYVE domain, it is unlikely that they can be
efficiently targeted to PtdIns3P substrate-containing
membranes.
The transcript levels of FAB1A and FAB1B were
investigated in a wide range of tissue types utilizing
publicly available microarray data with the GENEVESTIGATOR software (Zimmermann et al., 2004).
FAB1A and FAB1B transcripts were found in all plant
tissues to varying degrees (Supplemental Fig. S1).
FAB1A showed highest expression levels in pollen,
seed, and senescent leaves, whereas FAB1B transcript
levels were highest in the root hair zone, pollen, and
stamens. The microarray data reveal that FAB1A expression levels are significantly lower than FAB1B in
all tissue categories (Supplemental Fig. S1).
Isolation of FAB1A and FAB1B Knockout Lines
In order to obtain Arabidopsis lines with mutations
in FAB1A and FAB1B, SALK T-DNA lines carrying
insertions in these genes were identified. A number of
lines with T-DNA insertions in these genes were
identified. SALK_013923 and SALK_066673 were chosen for further investigation as they contained insertions in exons of FAB1A (At4g33240; exon 4 of 11) and
FAB1B (At3g14270; exon 9 of 11), respectively (Fig. 2A).
We predicted that these alleles ( fab1a-1 and fab1b-1)
would be unable to produce functional protein due to
the T-DNA insertions being positioned 5# of the lipid
kinase domain. PCR-based genotyping confirmed
T-DNA insertions in FAB1A and FAB1B. Homozygous
fab1a-1 and fab1b-1 mutant plant lines were generated
from these stocks and were checked for transcripts by
reverse transcription (RT)-PCR. FAB1A and FAB1B
transcripts could not be detected in fab1a-1/fab1a-1 or
fab1b-1/fab1b-1 plants, respectively, but were present
in wild-type plants (Supplemental Fig. S2). It should
be noted that the primers used to amplify cDNA by
RT-PCR were 3# of the T-DNA insertion sites; thus,
short transcripts coding for truncated proteins could
potentially be produced in the homozygous T-DNA
insertion lines. However, even if this were the case,
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Whitley et al.
Figure 1. Arabidopsis possesses two genes encoding FYVE domain-containing protein orthologs of PIKfyve/Fab1. A, Production
of PtdIns(3,5)P2 from PtdIns requires the sequential action of two enzymes, PtdIns 3-kinase (VPS34) and PtdIns3P 5-kinase
(PIKfyve/Fab1). FAB1 phosphorylates the D-5 position of the phosphoinoside ring. B and C, Arabidopsis has two genes, FAB1A
and FAB1B, encoding proteins with similar domain structure to PIKfyve/Fab1 proteins. PIKfyve/Fab1 proteins have been
identified in a broad range of species (C) and appear to be ubiquitous among eukaryotes. Arabidopsis has two genes, FAB1C and
FAB1D, encoding proteins that lack the PtdIns3P-binding FYVE domain (B). FYVE-domain-lacking proteins appear to be found
only among plant lineages (C). Species prefixes in C are as follows: Sp, Schizosaccharomyces pombe; Af, Aspergillus fumigatus;
Kl, Kluyveromyces lactis; Sc, Saccharomyces cerevisiae; Dd, Dictyostelium discoideum; Cr, Chlamydomonas reinhardtii; Os,
Oryza sativa; Vv, Vitis vinifera; At, Arabidopsis thaliana; Pt, Populus trichocarpa; Pp, Physcomitrella patens; Mt, Medicago
truncatula; Dm, Drosophila melanogaster; Dr, Danio rerio; Mm, Mus musculus; Hs, Homo sapiens.
truncated proteins would lack the C-terminal lipid
kinase domain and thus be unable to convert PtdIns3P
to PtdIns(3,5)P2. Therefore, we consider the fab1a-1/
fab1a-1 and fab1b-1/fab1b-1 plants to be null mutants of
FAB1A and FAB1B, respectively.
fab1a-1 and fab1b-1 Lines Display Subtle Leaf-Curling
Phenotypes and Are Reproductively Viable
Seeds from fab1a-1/fab1a-1 and fab1b-1/fab1b-1
plants exhibited normal germination rates and frequencies. Germination frequencies for both lines were
not significantly different from the wild type, being
greater than 95% on soil and greater than 98% on 13
Murashige and Skoog medium. Both mutant lines
appeared healthy and developed without displaying
any gross morphological defects. However, by 4 weeks
postgermination, it was noted that the leaves of both
fab1a-1/fab1a-1 and fab1b-1/fab1b-1 mutant lines appeared different from those of wild-type plants.
Leaves of mutant plants were noticeably curled at
their margins (Fig. 2B). A quantitative analysis of leaf
curling (Fig. 2C) revealed that there was a statistically
significant difference in the degree of curling of fab1b-1/
fab1b-1 compared with the wild type (P = 0.027).
Although there was a similar trend in the data for
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Role of PtdIns(3)P 5-Kinase in Pollen Development
fab1a-1/fab1a-1, the phenotype was not as severe (P =
0.066). Both fab1a-1/fab1a-1 and fab1b-1/fab1b-1 plant
lines were fully reproductively viable, displaying seed
set values indistinguishable from that of the wild type.
fab1a-1/fab1a-1;fab1b-1/fab1b-1 Double Mutants Cannot
Be Generated
Figure 2. Identification of a leaf curl phenotype in fab1a-1/fab1a-1 and
fab1b-1/fab1b-1 plants. A, Exon and intron structure of Arabidopsis
FAB1A and FAB1B genes. Boxes represent exons and intervening lines
represent introns. The locations and orientations of SALK T-DNA
insertions are indicated for both fab1a-1 and fab1b-1 alleles. LB and
RB indicate the left border and right border of the T-DNA, respectively.
Inward-facing arrows indicate the positions and directions of primers
used in RT-PCR experiments to verify that the T-DNA insertions
disrupted gene expression. The region of the genes encoding the
catalytic PtdIns3P 5-kinase domain are indicated (PIP5K). B, Plant lines
homozygous for fab1a-1 and fab1b-1 mutant alleles have a distinct leaf
curl phenotype in comparison with wild-type (WT) plants; plants
shown are 4 weeks old. C, The degree of curling was quantified by
comparing the ratio of the uncurled leaf area with the visible curled
surface area for each line. Leaf areas were determined by digital image
analysis using ImageJ software. Curling was most extreme in fab1b-1/
fab1b-1 plants, being significantly different from the wild type (P =
0.027). fab1a-1/fab1a-1 plants displayed a less extreme, although
consistent, phenotype in comparison with the wild type (P = 0.066).
Plotted are the ratio means with 95% confidence intervals indicated by
vertical bars. Leaves were analyzed from three randomly selected
plants from each line (numbers of leaves measured: wild type, n = 30;
fab1a-1/fab1a-1, n = 32; fab1b-1/fab1b-1, n = 32).
Due to the ubiquitous expression profile of FAB1A
and FAB1B and the lack of severe developmental
defects in mutant plant lines, it remains feasible that
FAB1A and FAB1B proteins are functionally redundant. In order to generate a double mutant (fab1a-1/
fab1a-1;fab1b-1/fab1b-1), both single mutant lines were
crossed with one another. The resulting F1 plants
(fab1a-1/+;fab1b-1/+) were allowed to self-pollinate,
and their progeny were screened by PCR for the
presence of double homozygous mutants. Of 86 plants
that were screened (Supplemental Table S1), not a
single double mutant was detected, despite there
being a theoretical one in 16 probability of recovering
such a plant (assuming no transmission defects). However, fab1a-1/+;fab1b-1/fab1b-1 and fab1a-1/fab1a-1;
fab1b-1/+ plants were identified, and these lines developed to maturity, again with no severe vegetative
morphological defects. This suggests that a single copy
of either FAB1A or FAB1B is sufficient to support
growth and development. In order to further increase
the chances of obtaining fab1a-1/fab1a-1;fab1b-1/fab1b-1
double mutants, fab1a-1/+;fab1b-1/fab1b-1 and fab1a1/fab1a-1;fab1b-1/+ plants were allowed to set self
seed. In these cases, potentially one-fourth of the
progeny should be homozygous for both mutant alleles, again assuming no transmission defects. Fifty
plants from each parent were screened by PCR to
determine their genotype, and again no double mutants were recovered.
Importantly, the seed set of both fab1a-1/+;fab1b-1/
fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants was normal (Fig. 3). As no gaps were observed in the developing siliques, this indicates that our inability to detect
double mutants was not due to failures in the ability of
plants to (1) produce female gametes or (2) undergo
seed development following fertilization. Furthermore, seeds harvested from these siliques germinated
Figure 3. Seed set in mature siliques of wild-type (WT), fab1a-1/+;
fab1b-1/fab1b-1, and fab1a-1/fab1a-1;fab1b-1/+ plant lines. fab1a-1/+;
fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants have seed set
indistinguishable from the wild type.
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Whitley et al.
with frequencies comparable to the wild type (99% on
13 Murashige and Skoog medium; n = 100). Taken
together, these data are strongly indicative of a transmission defect affecting only fab1a-1/fab1b-1 pollen.
fab1a-1/fab1b-1 Pollen Is Not Viable
Plants carrying three mutant alleles (fab1a-1/+;fab1b-1/
fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+) will produce
pollen of which 50% will lack both wild-type FAB1A
and FAB1B genes (i.e. will be of the genotype fab1a-1/
fab1b-1). Pollen derived from these plants was checked
for viability by staining with fluorescein diacetate
(FDA). A large proportion of pollen isolated from
fab1a-1/+;fab1b-1/fab1b-1 (34%) and fab1a-1/fab1a-1;
fab1b-1/+ (43%) plants failed to stain with FDA and
therefore was likely to be dead (Fig. 4; Table I). Many
of these grains were seen to be collapsed and were
frequently devoid of cytoplasmic content (Fig. 4B).
Staining of pollen with the nucleic acid stain 4#,6diamidino-phenylindole (DAPI) supported the FDA
data, as many grains lacked distinct sperm and vegetative cell nuclei. However, of those grains that
remained intact, none was found to have abnormalities in
the number of nuclei (two sperm nuclei and one
vegetative cell nucleus), suggesting that gametogenesis progresses normally for both wild-type and mutant
grains.
To further investigate whether fab1a-1/fab1b-1 pollen
is capable of effecting fertilization, restricted pollinations were carried out between fab1a-1/+;fab1b-1/
fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ pollen parents
and a transgenic male-sterile line (A9::barnase) as the
mother. Fifty progeny resulting from each of these
pollinations were screened for the presence of fab1a-1
or fab1b-1 mutant alleles, and none of the resulting
plants were found to have been sired by fab1a-1/
fab1b-1 pollen. Thus, we conclude that the failure to
generate fab1a-1/fab1a-1;fab1b-1/fab1b-1 double mutant plants is entirely attributable to fab1a-1/fab1b-1
pollen being inviable and not a case of it being at a
competitive disadvantage in pollinations of mixed
pollen genotype.
fab1a-1/fab1b-1 Pollen Displays an Abnormal Vacuolar
Phenotype Late in Pollen Development
To further investigate fab1a-1/fab1b-1 pollen inviability, fresh mature pollen, derived from fab1a-1/+;
fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants,
was stained with Neutral Red, which rapidly accumulates in vacuoles and vesicles. Shortly after pollen
hydration in the stain, approximately 50% of these
pollen grains had either an abnormal vacuolar phenotype or were unstained/dead (Table II; Fig. 4, F–I, K,
and L), whereas the vast majority of pollen grains from
wild-type plants (98%) displayed numerous small
vacuoles of relatively uniform size and morphology
(Table II; Fig. 4, E and J). The majority of grains that
were deemed abnormal were collapsed (Fig. 4, K and
Figure 4. Phenotypic analysis of mature pollen from wild-type, fab1a-1/+;
fab1b-1/fab1b-1, and fab1a-1/fab1a-1;fab1b-1/+ plants. A and B,
Fluorescence microscopy of FDA-stained pollen derived from wildtype (A) and fab1a-1/+;fab1b-1/fab1b-1 (B) plants. FDA stains live cells
green when imaged under UV light. A large proportion of pollen from
fab1a-1/+;fab1b-1/fab1b-1 plants was collapsed and failed to stain with
FDA (white arrowheads in B), indicating that they were dead. This was
also observed for pollen from fab1a-1/fab1a-1;fab1b-1/+ plants (data not
shown). C and D, DAPI staining of pollen nuclei revealed that uncollapsed
pollen grains from fab1a-1/+;fab1b-1/fab1b-1 plants (D) were trinucleate,
as they were for wild-type pollen (C). This was also observed for pollen
from fab1a-1/fab1a-1;fab1b-1/+ plants. White arrows indicate the diffuse
vegetative nucleus in close proximity to the more intensely fluorescing
pair of sperm nuclei. E to L, Neutral Red staining of pollen vacuoles observed by light differential interference contrast microscopy. Wild-type
pollen is characterized as containing many small, regular vacuoles (E
and J). Pollen samples from fab1a-1/fab1a-1;fab1b-1/+ (K) and fab1a-1/+;
fab1b-1/fab1b-1 (L) plants contained a large number of collapsed grains
(black arrowheads). A small proportion of the uncollapsed grains from
fab1a-1/+;fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants had a
range of highly abnormal vacuolar phenotypes, typified by the presence of relatively few, large, and frequently irregular vacuoles (F–I).
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Role of PtdIns(3)P 5-Kinase in Pollen Development
Table I. Analysis of pollen viability by FDA staining
The effect of fab1a-1 and fab1b-1 mutant alleles on pollen viability
was assessed by staining pollen from freshly dehisced anthers with
FDA as described in “Materials and Methods.” Viable grains stain FDA
positive (Fig. 4, A and B). Actual numbers of grains scored are
indicated in parentheses.
Genotype
Pollen FDA
Positive
Pollen FDA
Negative
%
%
Wild type
fab1a-1/+;fab1b-1/fab1b-1
fab1a-1/fab1a-1;fab1b-1/+
99 (198)
66 (145)
57 (166)
1 (2)
34 (75)
43 (125)
L), although some remained expanded and displayed
a range of phenotypes typified by the presence of
relatively few vacuoles that were irregular in size and
morphology, with some vacuoles being exceedingly
large (Fig. 4, F–I).
In order to assess at which stage of pollen development the fab1a-1/fab1b-1 mutations exerted their effects, chemically fixed floral tissues from fab1a-1/+;
fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants
were sectioned and observations were made with a
light microscope. No evidence of aberrant pollen/
vacuolar morphology could be detected at early developmental stages, with tetrads and the highly vacuolated uninucleate microspores looking uniform and
identical to pollen in anthers from wild-type plants
(data not shown). Uninucleate microspores then divide to form bicellular pollen consisting of a large
vegetative cell, which fills the pollen wall cavity, and
the generative cell, which resides within the vegetative
cell cytoplasm. The transition from the uninucleate to
the bicellular stage of development is followed immediately by fragmentation of the large vacuole that
predominated in uninucleate microspores (McCormick,
1993; Yamamoto et al., 2003). We observed no significant differences between wild-type pollen and pollen
from fab1a-1/+;fab1b-1/fab1b-1 and fab1a-1/fab1a-1;
fab1b-1/+ plants at this stage of development. As
development proceeded toward the second mitotic
division, which produces two sperm cells from the
generative cell, there was some indication that vacuoles were enlarged and irregularly sized in a proportion of grains from the two mutant lines compared
with the wild type (Fig. 5, A, C, and E). This abnormal
vacuolar phenotype became more marked just prior to
and after the second mitotic division in both mutant
lines (Fig. 5, G and H), whereas vacuoles of wild-type
grains appeared to fragment into many smaller vacuoles and disperse at the transition from the bicellular
to the tricellular stage of development (Yamamoto
et al., 2003). As tricellular pollen matured in wild-type
anthers, the pollen cytoplasm took on a smoother
appearance, with vacuoles being difficult to detect
with certainty among many other membrane-bound
structures. In contrast, nearly 50% of grains from
mutant lines underwent rapid collapse or exhibited
extreme vacuolar morphologies as tricellular pollen
matured (Fig. 5, D and F). Taken together, these
observations strongly suggest that fab1a-1/fab1b-1 pollen grains develop normally until just after the first
mitotic division. However, following this, during the
phase of extensive vacuole fragmentation that occurs
prior to the second mitotic division, these grains fail to
regulate this process properly.
Further analysis of pollen development by transmission electron microscopy for fab1a-1/+;fab1b-1/
fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants confirmed
the observations obtained by light microscopy. The
ultrastructure of “early” bicellular pollen was highly
similar between wild-type (Fig. 6, A and B) and
mutant (Fig. 6, E, F, I, and J) lines. However, highly
aberrant vacuolar phenotypes were observed in grains
from mutant plant lines at the tricellular stage of
development (Fig. 6, G, H, K, and L). These vacuoles
were frequently very large and irregular in shape with
uneven boundaries (Fig. 6, G and H), starkly contrasting with the small, more uniform vacuoles with
smooth peripheries present in wild-type grains (Fig.
6, C and D). Many of these large vacuoles contained
granular deposits that appeared to be degraded cytoplasm. In many cases, these vacuoles filled the majority of the pollen cavity, suggesting large-scale loss of
cytoplasm and degradation of cellular components.
Again, many grains were completely collapsed and
contained little or no cytoplasm (Fig. 6K).
Table II. Assessment of pollen vacuolar phenotype by Neutral Red staining
The effect of fab1a-1 and fab1b-1 mutant alleles on the morphology of vacuoles in pollen collected
from freshly dehisced anthers was assessed by staining with Neutral Red as described in “Materials and
Methods.” Pollen scored as “normal” contained numerous small vacuoles (Fig. 4E), whereas grains scored
as “abnormal” typically contained fewer irregularly sized large vacuoles (Fig. 4, F–I) or were collapsed.
“Unstained” pollen grains lacked cytoplasm and were frequently collapsed. Actual numbers of grains
scored are indicated in parentheses.
Genotype
Wild type
fab1a-1/+;fab1b-1/fab1b-1
fab1a-1/fab1a-1;fab1b-1/+
Normal Pollen
Abnormal Pollen
%
%
Unstained Pollen
%
98 (191)
55.4 (210)
48.1 (140)
1.5 (3)
30.1 (114)
19.6 (57)
0.5 (1)
14.5 (55)
32.3 (94)
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Whitley et al.
DISCUSSION
Figure 5. Phenotypic analysis by light microscopy of pollen development in wild-type, fab1a-1/fab1a-1;fab1b-1/+, and fab1a-1/+;fab1b-1/
fab1b-1 plant lines. Pollen grains in anthers at bicellular and tricellular
stages of development are shown, respectively, for wild-type (A and B),
fab1a-1/fab1a-1;fab1b-1/+ (C and D), and fab1a-1/+;fab1b-1/fab1b-1
(E and F) plant lines. Wild-type pollen displayed multiple small
vacuoles at both bicellular and tricellular stages of development (A
and B). Some bicellular pollen grains in mutant lines contained abnormally large vacuoles (black arrowheads in C and E). A large proportion of
tricellular grains in mutant lines were collapsed (black stars in D and F),
although some were only partially collapsed and contained very large
vacuoles (black arrowheads in F). The abnormal vacuolar phenotype
was noted to be more marked in bicellular grains approaching the
second mitotic division in both fab1a-1/fab1a-1;fab1b-1/+ (G) and
fab1a-1/+;fab1b-1/fab1b-1 (H) plant lines (black arrowheads indicate
PtdIns(3,5)P2 is a ubiquitous but minor membrane
phospholipid in plants, animals, and yeast (Dove et al.,
1997; Whiteford et al., 1997; Meijer et al., 1999). It has a
conserved role in regulating endomembrane trafficking events in animals and yeast (for review, see Dove
et al., 2009). PtdIns(3,5)P2 production is tightly regulated (Sbrissa et al., 2007; Botelho et al., 2008; Ikonomov
et al., 2009) and inducible by a number of different
stimuli. For example, hyperosmotic stress increases
levels of PtdIns(3,5)P2 by over 20-fold in yeast (Dove
et al., 1997) and significantly increases levels in animal
cells and plants (Meijer et al., 1999; Zonia and Munnik,
2004; Sbrissa and Shisheva, 2005). Hormonal stimuli,
such as insulin in mammalian cells, can also lead to an
increased production of PtdIns(3,5)P2 (Ikonomov et al.,
2007).
In animals and yeast, PtdIns(3,5)P2 is produced from
PtdIns3P by the action of PtdIns3P 5-kinases called
PIKfyve and Fab1p, respectively (Shisheva, 2008).
PIKfyve and Fab1p have similar domain architecture,
consisting of an N-terminal FYVE domain that recruits
the protein to specific membranes by binding PtdIns3P
(Sbrissa et al., 2002), a central Cpn60_TCP1 domain
that probably plays a regulatory role, and a kinase
domain that is responsible for the conversion of
PtdIns3P to PtdIns(3,5)P2. In Arabidopsis, the two
genes encoding proteins with this domain architecture
are FAB1A (At4g33240) and FAB1B (At3g14270). In
addition, there are two genes, FAB1C (At1g71010) and
FAB1D (At1g34260), that contain a Cpn60_TCP1 domain and a C-terminal lipid kinase domain but lack a
FYVE domain. As the FYVE domain is not essential for
yeast Fab1p to phenotypically rescue a fab1D strain
(Botelho et al., 2008), the possibility that FAB1C and
FAB1D are PtdIns3P 5-kinases cannot be ruled out.
The only obvious abnormal feature of FAB1A and
FAB1B homozygous T-DNA insertion lines was a leafcurling phenotype. Given the apparent evolutionarily
conserved role of PtdIns(3,5)P2 in osmoregulation, the
inability of fab1a-1/fab1a-1 and fab1b-1/fab1b-1 lines to
produce sufficient levels of this lipid to correctly
regulate turgor pressure may explain the leaf curling.
The severity of leaf curl was most pronounced in fab1b-1/
fab1b-1 plants, consistent with FAB1B being more
highly expressed than FAB1A in leaves (Supplemental
Fig. S1). Despite phenotypic differences between both
the homozygous single mutant lines and wild-type
plants, the mutants developed to maturity and exhibited normal seed set. The broadly overlapping
expression profiles of FAB1A and FAB1B, together
with similar phenotypes being exhibited by fab1a-1/
fab1a-1 and fab1b-1/fab1b-1 mutants, suggest that
FAB1A and FAB1B are, to a large extent, functionally
redundant. We were unable to assess developmental
abnormally large vacuoles). GC, Generative cell; SC, sperm cell; Tp,
tapetum; VN, vegetative nucleus. Bars = 20 mm.
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Role of PtdIns(3)P 5-Kinase in Pollen Development
Figure 6. Transmission electron microscopy of pollen from wild-type,
fab1a-1/+;fab1b-1/fab1b-1, and fab1a-1
/fab1a-1;fab1b-1/+ plants. Pollen was
observed at bicellular (wild-type,
A and B; fab1a-1/fab1a-1;fab1b-1/+,
E and F; fab1a-1/+;fab1b-1/fab1b-1,
I and J) and tricellular (wild-type, C
and D; fab1a-1/fab1a-1;fab1b-1/+, G
and H; fab1a-1/+;fab1b-1/fab1b-1, K
and L) stages of development. Dashed
boxes indicate enlarged regions in adjacent micrographs to the right. Many
small vacuoles are present in bicellular
pollen (white arrowheads in B, F, and
J), although some abnormally large
vacuoles were observed in a proportion of grains from mutant lines (I and
J). Wild-type tricellular grains were
characterized by the presence of
many small regular vacuoles (white
arrowheads in D), whereas approximately 50% of tricellular grains from
both mutant lines were either collapsed or contained large irregular
vacuoles (black stars in G, H, K, and
L) in addition to enlarged regular
vacuoles (white arrowhead in H). Ex,
Exine; GC, generative cell; It, intine;
LB, lipid body. Bars = 5 mm.
phenotypes in fab1a-1/fab1a-1;fab1b-1/fab1b-1 double
mutant plants, as attempts to generate such mutants
were unsuccessful. Genetic analyses revealed unambiguously that fab1a-1/fab1b-1 pollen was incapable of
transmitting these alleles to the next generation. If the
role of FAB1A and FAB1B in growth and development
is to be fully elucidated, transgenic approaches aimed
at abolishing their respective transcripts will have to
be employed.
Importantly, it has been previously reported that
Arabidopsis pollen lacking the PtdIns 3-kinase VPS34
have severe developmental defects (Lee et al., 2008b).
VPS34 produces PtdIns3P from PtdIns; as PtdIns3P is
a substrate for Fab1/PIKfyve enzymes, and therefore
required for PtdIns(3,5)P2 synthesis, the defects seen in
VPS34-deficient pollen could be primarily due to a
lack of PtdIns(3,5)P2 production (rather than an absence of PtdIns3P). In order to test this hypothesis,
pollen development in plants carrying fab1a-1 and
fab1b-1 mutant alleles was investigated. In fab1a-1/+;
fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants,
50% of the pollen will have a fab1a-1/fab1b-1 genotype.
A large proportion of pollen from these plants did not
stain with FDA and therefore was considered to be
nonviable/dead pollen (34% and 43%, respectively).
We predict that the dead pollen is mainly fab1a-1/
fab1b-1 in genotype, as pollen from fab1a-1/fab1a-1
(fab1a-1/FAB1B pollen) and fab1b-1/fab1b-1 (FAB1A/
fab1b-1 pollen) plants appear normal (.98% FDA
positive). Although some fab1a-1/fab1b-1 pollen sur-
vives to maturity, Neutral Red staining revealed a
further population of grains with aberrant vacuolar
morphologies. Together, these grains and those that
were observed to be dead account for the 50% of fab1a-1/
fab1b-1 pollen in the population. Another notable feature of pollen from fab1a-1/+;fab1b-1/fab1b-1 and
fab1a-1/fab1a-1;fab1b-1/+ plants was that it tended to
aggregate and was difficult to remove from anthers.
No such problems were noted for pollen derived from
either fab1a-1/fab1a-1 or fab1b-1/fab1b-1 plants.
The phenotypes observed in pollen from fab1a-1/+;
fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants
are very similar to those described for VPS34 mutant
plants (Lee et al., 2008b). However, there are some
important differences. Some transmission of the vps34-1
allele was observed by Lee et al. (2008b), but despite
extensive efforts, transmission of fab1a-1/fab1b-1 pollen
was never observed in our study. In addition to the
abnormal vacuolar phenotype, vps34 mutant plants
produce an increased proportion of binucleate pollen
(Lee et al., 2008b). It was proposed that abnormal
vacuoles could physically prevent normal cell division
in developing mutant pollen or alternatively that
production of PtdIns3P may have a more specific
role in directly regulating the cell cycle. As fab1a-1/
fab1b-1 pollen has abnormal vacuoles but no cell
division defect, we favor the latter of these explanations. We propose that the vacuolar defects observed
in pollen from vps34-1 mutant plants and in fab1a-1/
fab1b-1 pollen can be entirely attributable to the inabil-
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Whitley et al.
ity of mutant pollen to produce normal levels of
PtdIns(3,5)P2.
An important question to consider is how does the
lack of FAB1A/FAB1B affect pollen viability? One
possibility is that PtdIns(3,5)P2 production may be
required for pollen tube growth, although, as a large
proportion of developing pollen in anthers of fab1a-1/
+;fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants
collapses long before dehiscence, it is apparent that the
primary defect in pollen occurs while the pollen is still
in the anthers. However, as it is known that membrane
dynamics are important for polarized pollen tube
elongation and that phosphoinositides play a key
role in membrane trafficking and cytoskeletal reorganization during polarized cell growth (Monteiro et al.,
2005; Samaj et al., 2006; Zonia and Munnik, 2008),
PtdIns(3,5)P2 may have such a role in surviving
fab1a-1/fab1b-1 pollen. Experimental evidence to address the possibility of FAB1A and FAB1B being
required for pollen tube growth was not obtained, as
our data suggest that only a small proportion of pollen
surviving beyond dehiscence was fab1a-1/fab1b-1,
with the majority of surviving pollen being fab1a-1/
FAB1B or FABlA/fab1b-1 (dependent on parent).
The presence of large vacuolar structures at the late
bicellular stage is the first sign that pollen development is perturbed in mutant plants. An increase in
PtdIns(3,5)P2 levels increases the surface area-tovolume ratio of intracellular organelles by mediating
fission of large vacuoles (or equivalent organelles)
into smaller vacuoles, protecting cells against rapid
dehydration and lysis (Weisman, 2003; Sbrissa and
Shisheva, 2005). In fab1a-1/fab1b-1 pollen, unable to
produce normal levels of PtdIns(3,5)P2, an inability of
vacuoles to undergo fission can explain the accumulation of large vacuoles at the late bicellular stage of
development. This would ultimately lead to pollen
collapse during the dehydration phase that occurs
just prior to anthesis. The cue for vacuole fission
could be strictly developmentally regulated or induced
by changes in solute concentration late in pollen development.
We are aware that the interpretation of these results
relies on the assumption that Arabidopsis FAB1A and
FAB1B are major sources of PtdIns(3,5)P2 production,
particularly in pollen. Due to the domain structure, we
feel that we can safely assume that FAB1A and FAB1B
will be PtdIns3P 5-kinases. However, it will be extremely difficult to biochemically quantify PtdIns(3,5)P2
levels with accuracy in fab1a-1/fab1b-1 pollen by metabolic labeling methods (Zonia and Munnik, 2004), due to
the high mortality of fab1a-1/fab1b-1 pollen among other
genotypes from fab1a-1/+;fab1b-1/fab1b-1 and fab1a-1/
fab1a-1;fab1b-1/+ plants.
Regarding the importance of FAB1A and FAB1B in
plant growth and development, microarray data reveal that their expression levels are very high in pollen
in comparison with most other tissues. This contrasts
with FAB1C expression, which is negligible in pollen.
Although FAB1D levels are relatively high in pollen,
this seems likely to be due to extremely high expression in sperm cells (Supplemental Fig. S1). Interestingly, lack of FAB1A and FAB1B in the female
gametophyte did not appear to have any measurable
effect on fertility. Thus, it is not gametogenesis per se
that requires FAB1A and FAB1B; rather, their function
in reproduction is related to specific developmental
processes unique to pollen. We speculate that this may
be related to preparation for the dehydration of pollen
prior to dehiscence, whereas no such stresses are
developmentally imposed within the ovule.
In summary, we present, to our knowledge, the first
genetic and phenotypic analysis of Fab1/PIKfyve
family proteins in plants. We conclude that the activity
of FAB1 proteins is essential for the development of
viable pollen. Our results are consistent with a role for
FAB1 proteins in responding to osmotic changes by
controlling levels of PtdIns(3,5)P2. This is most apparent in pollen, but the leaf curl phenotype in homozygous single mutant plants further hints at a defect in
osmoregulation. Future studies will include investigations of cell and plant phenotypes grown under
normal and stress conditions in conjunction with
biochemical analysis of PtdIns(3,5)P2 levels in mutant
plants. In the future, it will be interesting to address
whether plant orthologs of known yeast and animal
effectors of PtdIns(3,5)P2 are essential for pollen viability or whether plant-specific effectors are responsible.
MATERIALS AND METHODS
Plant Growth Conditions
All Arabidopsis (Arabidopsis thaliana) lines used in this study were grown
in a controlled-environment room with a 16-h-day/8-h-night cycle. Temperature was maintained at 22°C 6 1°C, and relative humidity was 60%. Seeds
were sown in petri dishes containing a fine-grade peat-based compost, and
seedlings were transferred to pots containing the same compost 5 d after
germination.
Phylogenetic Analysis of the FAB1/PIKfyve Family
of Proteins
Protein sequences were aligned using ClustalX (Larkin et al., 2007), and
trees were drawn using the NJPLOT program distributed with ClustalX
software. The following UniProtKB protein sequences were used for the
analysis: Schizosaccharomyces pombe, FAB1_SCHPO; Aspergillus fumigatus,
Q4WN65_ASPFU; Kluyveromyces lactis, Q6CS22_KLULA; Saccharomyces cerevisiae, FAB1_YEAST; Dictyostelium discoideum, B0G126_DICDI; Chlamydomonas
reinhardtii, A8J5R5_CHLRE; Oryza sativa, Q6ZJN4_ORYSJ, Q8H3L8_ORYSJ,
and Q0DRB6_ORYSJ; Vitis vinifera, A5B6P1_VITVI, A7PDP8_VITVI, and
A5C675_VITVI; Arabidopsis, Q0WUR5_ARATH (FAB1A), Q9LUM0_ARATH
(FAB1B), Q9SSJ8_ARATH (FAB1C), and Q9XID0_ARATH (FAB1D);
Populus trichocarpa, B9HJA2_POPTR and B9N7I0_POPTR; Physcomitrella
patens, A9SCF9_PHYPA and A9RYY6_PHYPA; Medicago truncatula,
A4Q7Q7_MEDTR; Drosophila melanogaster, FYV1_DROME; Danio rerio,
B2KTE1_DANRE; Mus musculus, FYV1_MOUSE; and Homo sapiens, FYV1_
HUMAN.
Isolation of FAB1A and FAB1B Mutant Plant Lines
SALK T-DNA insertion lines (Alonso et al., 2003) were obtained from the
Nottingham Arabidopsis Stock Centre. Genotyping of plants was carried out
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Role of PtdIns(3)P 5-Kinase in Pollen Development
Supplemental Figure S1. Expression profiles of Arabidopsis FAB1A,
FAB1B, FAB1C, and FAB1D.
by PCR using genomic DNA as a template. fab1a-1 alleles (SALK_013923) were
identified using the following primers: T-DNA-specific left border primer
(5#-ATTTTGCCGATTTCGGAAC-3#) and gene-specific left (LP; 5#-CCTGTCGTTTCCATCCAATTT-3#) and right (RP; 5#-TGGACATGAATCCTTCGACTG-3#) primers. fab1b-1 alleles (SALK_066673) were identified using the
same left border primer in conjunction with gene-specific LP (5#-TGAACCATTGCTTCTGCTAGG-3#) and RP (5#-CTCTCGTGGACATTCTTGAGC-3#)
primers. RT-PCR was carried out to test for the presence of transcripts in
fab1a-1/fab1a-1 plants using LP (5#-CCAGGCAGCTATCATTTCACG-3#) and
RP (5#-GAAAGCCATTGATCCGAGGAC-3#) primers. Likewise fab1b-1/
fab1b-1 plants were also tested by RT-PCR using LP (5#-TGGTTCAGTCGACGAGAGCAT-3#) and RP (5#-GGAGACCATTGATCAGGGACC-3#) primers.
Total RNA for RT-PCR was isolated using the RNeasy kit (Qiagen) with the
manufacturer’s instructions being followed.
fab1a-1/+;fab1b-1/fab1b-1 and fab1a-1/fab1a-1;fab1b-1/+ plants were isolated by initially crossing fab1a-1/fab1a-1 and fab1b-1/fab1b-1 single mutant
lines to generate F1 seeds. The F1 plants were allowed to set “self” seed to give
an F2 population from which the lines were recovered.
We thank Dr. Alex Jeffries for guidance with the phylogenetic analysis,
Ursula Potter for technical assistance with electron microscopy, Dr. Baoxiu Qi
for academic and technical input, and Jiannan Guo for initial PCR characterization of plant lines. We also thank the Salk Institute Genomic Analysis
Laboratory and the Nottingham Arabidopsis Stock Centre for provision of
mutant seed.
Leaf Curl Analysis
Received August 13, 2009; accepted October 18, 2009; published October 21,
2009.
The degree of leaf curl was quantified for wild-type, fab1a-1/fab1a-1 and
fab1b-1/fab1b-1 plants by estimating the visible leaf area in photographs taken
directly above plants and comparing this area with actual leaf area. Leaves
were removed sequentially from the plants to permit a full analysis. Each leaf
was then flattened under clear adhesive plastic and rephotographed. Image
analysis was carried out using ImageJ software (Abramoff et al., 2004) to
obtain accurate measurements.
Pollen Analysis
For all staining methods, pollen was initially collected from six open
flowers placed in 400 mL of an 8% (w/v) Suc solution. The flowers were briefly
vortexed to release pollen and then removed with tweezers. Pollen grains
were pelleted by centrifugation at 6,000g for 2 min and then resuspended in
the appropriate stain for microscopical analysis.
Viability of pollen grains was assessed by staining with FDA. Pollen was
immersed in 8% (w/v) Suc containing 0.5 mg mL21 FDA and observed
immediately. Pollen nuclei were stained with DAPI as described by Lee et al.
(2008b), except that 8% (w/v) Suc was added to the staining solution. Vacuoles
were visualized in pollen by staining with 0.02% (w/v) Neutral Red in 8% (w/v)
Suc. Observations were carried out with a Nikon ECLIPSE 90i microscope
(Nikon Instruments) equipped with epifluorescence modules for UV microscopy.
Analysis of Pollen Development by Light and
Transmission Electron Microscopy
Material from wild-type, fab1a-1/+;fab1b-1/fab1b-1, and fab1a-1/fab1a-1;
fab1b-1/+ plants was prepared using a method adapted from that described by
Lee et al. (2008b). Buds and anthers were fixed in a solution containing 2.5%
glutaraldehyde and 4% paraformaldehyde in a 0.1 M PIPES buffer (pH 7.4) at
4°C overnight. Samples were then rinsed in PIPES buffer. Postfixation was
carried out in 1% (w/v) osmium tetroxide for 4 h at 4°C, following which the
material was washed in distilled water. Samples were stained in 1% aqueous
uranyl acetate for 1 h in the dark and then dehydrated in an ethanol series
(30%, 50%, 70%, 80%, 90%, 95%, and 100%) and embedded in LR White resin
(London Resin Company). Sections (0.5 mm thick) were cut and stained with
Toluidine Blue for light microscopy. Sections (100 nm thick) were cut for
transmission electron microscopy. Light microscopy was conducted with a
Nikon ECLIPSE 90i microscope (Nikon Instruments). For transmission electron microscopy, observations were made with a JEOL 1200EXII transmission
electron microscope operating at 80 kV.
Sequence data from this article can be found in the GenBank/EMBL data
libraries under accession numbers NM_119478 and NM_112285.
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure S2. Expression of FAB1A and FAB1B in wild-type
and mutant Arabidopsis plants.
Supplemental Table S1. Expected and observed genotypes of the progeny
derived from self-pollinated fab1a-1/FAB1A;fab1b-1/FAB1B plants.
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