Download γ-Tubulin Is Essential for Microtubule Organization and

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g-Tubulin Is Essential for Microtubule Organization
and Development in Arabidopsis
W
Martine Pastuglia,a,1,2 Juliette Azimzadeh,a,1,3 Magali Goussot,a Christine Camilleri,a Katia Belcram,a
Jean-Luc Evrard,b Anne-Catherine Schmit,b Philippe Guerche,a and David Boucheza
a Station de Génétique et d’Amélioration des Plantes, Institut National de la Recherche Agronomique,
78026 Versailles Cedex, France
b Institut de Biologie Moléculaire des Plantes, Centre National de la Recherche Scientifique, 67084 Strasbourg Cedex, France
The process of microtubule nucleation in plant cells is still a major question in plant cell biology. g-Tubulin is known as one
of the key molecular players for microtubule nucleation in animal and fungal cells. Here, we provide genetic evidence that in
Arabidopsis thaliana, g-tubulin is required for the formation of spindle, phragmoplast, and cortical microtubule arrays. We
used a reverse genetics approach to investigate the role of the two Arabidopsis g-tubulin genes in plant development and in
the formation of microtubule arrays. Isolation of mutants in each gene and analysis of two combinations of g-tubulin double
mutants showed that the two genes have redundant functions. The first combination is lethal at the gametophytic stage.
Disruption of both g-tubulin genes causes aberrant spindle and phragmoplast structures and alters nuclear division in
gametophytes. The second combination of g-tubulin alleles affects late seedling development, ultimately leading to lethality
3 weeks after germination. This partially viable mutant combination enabled us to follow dynamically the effects of g-tubulin
depletion on microtubule arrays in dividing cells using a green fluorescent protein marker. These results establish the
central role of g-tubulin in the formation and organization of microtubule arrays in Arabidopsis.
INTRODUCTION
Microtubules are highly dynamic polar polymers of noncovalently bound ab-tubulin heterodimers that are major structural
components of the cytoskeleton in eukaryotic cells. In animal and
fungal cells, microtubule nucleation takes place at conspicuous
microtubule-organizing centers (MTOCs), such as the centrosome or the spindle pole body, whose activity determines the
spatial and temporal organization of the microtubule cytoskeleton. Higher plant cells lack discrete MTOCs but assemble highly
ordered arrays of microtubules that coordinate cell division and
expansion (Wasteneys, 2002). The preprophase band forms
during late G2 at the cell cortex and delineates the future site of
division. It is replaced during late prophase by an acentriolar
mitotic spindle. The phragmoplast, assembled during late anaphase, promotes the synthesis of the new cell plate separating
daughter cells. As they enter G1, a cortical microtubule array is
formed at the cell cortex and participates in the control of cell
elongation and cell wall deposition.
1 These
authors contributed equally to this work.
whom correspondence should be addressed. E-mail pastugli@
versailles.inra.fr; fax 33-130-83-3319.
3 Current address: Institut Curie, Centre National de la Recherche
Scientifique/Unité Mixte de Recherche 144, 26 rue d’Ulm, 75005 Paris,
France.
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.plantcell.org) is: Martine Pastuglia
([email protected]).
W
Online version contains Web-only data.
Article, publication date, and citation information can be found at
www.plantcell.org/cgi/doi/10.1105/tpc.105.039644.
2 To
Different microtubule nucleation sites accounting for the assembly of plant microtubule arrays have been established in
plant cells. The nuclear envelope has been shown to retain
nucleating activity in vitro (Stoppin et al., 1994). Using fluorescenttagged microtubule markers, in vivo nucleation has been revealed at the cell cortex, either at sites linked to preexisting
microtubules (Shaw et al., 2003; Van Bruaene et al., 2004; Murata
et al., 2005) or de novo at sites with no other detectable microtubules (Shaw et al., 2003). Fluorescent-tagged protein markers,
such as ATEB1A–green fluorescent protein (GFP), have also
provided support for the existence of diffuse, mobile nucleation
sites at spindle poles (Chan et al., 2003). However, our knowledge about the molecular composition of plant microtubule
nucleating sites is still in its infancy. In animals and fungi, a large
body of evidence strongly implicates the highly conserved
g-tubulin protein, the third member of the tubulin protein family,
as a key element for microtubule nucleation at MTOCs (reviewed
in Jeng and Stearns, 1999; Moritz and Agard, 2001; Job et al.,
2003; Jaspersen and Winey, 2004). Apart from the g-tubulin pool
present at the centrosome, most soluble cytoplasmic g-tubulin
is part of large complexes named g-tubulin ring complexes
(g-TuRCs) containing 10 to 13 g-tubulin molecules per complex
and at least eight proteins in addition to g-tubulin. g-Tubulin
present at the centrosome likely comes from association of
g-TuRCs with the pericentriolar matrix. Animal g-TuRC is seen as
an open ring of the approximate diameter of a microtubule that
caps the minus end of a microtubule, but the precise mechanism
of microtubule nucleation by g-TuRCs is still being debated.
Smaller complexes called g-tubulin small complexes have
also been found in many cell types and are clearly a component
of g-TuRCs. They consist of two g-tubulin molecules and one
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The Plant Cell
molecule each of two related proteins known as Spc97 and
Spc98 proteins in Saccharomyces cerevisiae.
In plants, the role of g-tubulin is still a debated question. Plant
g-tubulin is present in protein complexes of various sizes in
maize (Zea mays), Arabidopsis thaliana, and fava bean (Vicia
faba) (Stoppin-Mellet et al., 2000; Drykova et al., 2003), and large
g-tubulin–containing complexes have nucleation activity in Arabidopsis (Drykova et al., 2003). The ability of plant g-tubulin to
nucleate microtubules was demonstrated by heterologous expression of Arabidopsis g-tubulin in fission yeast lacking endogenous g-tubulin. Arabidopsis g-tubulin was able to bind MTOCs
and nucleate microtubule assembly in Schizosaccharomyces
pombe (Horio and Oakley, 2003). Another piece of evidence
favoring a role of g-tubulin in microtubule nucleation comes from
a study on the subcellular localization of the plant Spc98 homologue. This protein, which was recently shown to be required for
microtubule nucleation on isolated plant nuclei, colocalized with
g-tubulin on the nuclear surface (Erhardt et al., 2002). However,
the unusual subcellular localization of g-tubulin in plant cells,
which seemed incompatible with a mere role in nucleation,
puzzled cell biologists for long. Indeed, in addition to its presence
at established nucleation sites, such as the nuclear surface in
Arabidopsis, wheat (Triticum aestivum), soybean (Glycine max),
and BY2 cells (Liu et al., 1993, 1994; Joshi and Palevitz, 1996;
Erhardt et al., 2002), or the acentriolar polar organizer of basal
land plants (Brown et al., 2004; Shimamura et al., 2004), plant
g-tubulin associates to all microtubule arrays along microtubules
in a punctuate manner and is not restricted to microtubule minus
ends, which would be expected for a protein supposedly involved in nucleation (Liu et al., 1993, 1994; Joshi and Palevitz,
1996; Panteris et al., 2000). Some aspects of this unusual localization have been clarified recently: Using a cell-free system,
Murata et al. (2005) demonstrated that this punctuate labeling on
cortical microtubules represents bona fide sites of nucleation on
the sides of extant microtubules, resulting in branched structures
(Murata et al., 2005). They also showed that g-tubulin is required
for this process. Although, in such a cell-free system many
parameters (e.g., local tubulin concentrations) may differ markedly from in vivo conditions, these results provide direct evidence of the involvement of g-tubulin in nucleation of cortical
microtubules. Whether, quantitatively speaking, this process
represents a major mechanism in vivo still needs to be evaluated
in addition to its involvement in the formation of mitotic arrays. In
particular, the fact that the Arabidopsis Spc98 homologue, although present at the cortex, is not codistributed with g-tubulin
on the whole length of microtubules needs to be clarified (Erhardt
et al., 2002).
Apart from its key role in microtubule nucleation, additional
functions for g-tubulin have been suggested in several eukaryotic systems. Recent genetic and molecular studies in fission
and budding yeast, in Aspergillus nidulans, and in Drosophila
melanogaster have revealed that g-tubulin could be involved
in microtubule dynamics or organization (Paluh et al., 2000;
Jung et al., 2001; Vogel et al., 2001) and in the control of
mitotic checkpoint and coordination of late mitotic events
(Hendrickson et al., 2001; Sampaio et al., 2001; Prigozhina
et al., 2004). Evidence for such roles is needed for plant
g-tubulin.
To gain further insights into the function of g-tubulin in plant
cells, we have obtained T-DNA insertion mutant lines for the two
genes encoding g-tubulin in Arabidopsis (Liu et al., 1994). Here,
we show that the two Arabidopsis genes are functionally redundant. We studied two double mutant combinations of TUBG1
and TUBG2 insertion alleles and investigated the effect of
g-tubulin depletion on plant development and organization of
mitotic and interphase microtubules arrays. Our results demonstrate the in vivo role of g-tubulin in the organization of all microtubule structures in plant cells, both during interphase and
cell division.
RESULTS
Isolation and Characterization of Arabidopsis
g-Tubulin Mutants
The complete sequence of the Arabidopsis genome confirmed
the occurrence of two highly similar genes encoding bona fide
g-tubulin isoforms, TUBG1 (At3g61650) and TUBG2 (At5g05620)
(Liu et al., 1994; Arabidopsis Genome Initiative, 2000), which
share 98% protein sequence identity. RT-PCR analysis using
gene-specific primers indicated that both g-tubulin genes are
constitutively expressed at high levels in all organs tested (Figure
1C). Affymetrix ATH1 data, corresponding to the combined expression of both g-tubulin genes (probe set 251331_s_at), confirms that global transcript level is high and rather constant during
development, although especially high in the shoot apical meristem and in cultured cells, and very low in pollen (Zimmermann
et al., 2004). To obtain mutants for both genes, we used either
PCR screening of a T-DNA–mutagenized Arabidopsis population
(Bechtold et al., 1993) or in silico searching in public T-DNA
databases (Alonso et al., 2003). We identified one TUBG1 and
two TUBG2 insertion lines: tubg1-1 harbors a T-DNA insert in the
first exon of TUBG1 associated with a 55-bp deletion of the
coding region, and tubg2-1 carries an insertion and a large deletion of most of the TUBG2 gene, whereas tubg2-2 has a
T-DNA insert in the 59 untranslated region of TUBG2 (Figures
1A and 1B). DNA sequencing of the T-DNA flanking regions and
DNA gel blot analysis enabled us to characterize the three insertion loci at the molecular level (Figures 1A and 1B).
RT-PCR with gene-specific primers indicated that no wildtype TUBG1 transcript is detectable in tubg1-1 mutants (Figure
1C). As often observed in T-DNA insertion lines, the 35S promoter present in the T-DNA produces fusion transcripts with
adjacent genomic regions: this is observed both in tubg1-1 and
tubg2-2, which originate from different T-DNA populations (Figure 1C). tubg2-1 is fully deleted for the TUBG2 coding sequence.
All three homozygous single mutant lines had a wild-type
phenotype in terms of growth and development and were fully
fertile. In order to study the effects of a simultaneous deficiency
for both g-tubulin isoforms, we crossed homozygous tubg1-1
plants with either tubg2-1 or tubg2-2 homozygous plants. In both
cases, neither double heterozygote F1 progenies nor homozygote/heterozygote F2 plants showed any vegetative defects,
showing that one functional copy of g-tubulin (out of four in
the wild type) is enough to sustain growth and development of
plants in standard conditions. However, the tubg1-1 tubg2-1
g-Tubulin Function in Arabidopsis
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Figure 1. Isolation of T-DNA Insertions in the Two g-Tubulin Genes and Protein Gel Blot Analysis of Double Mutant Plants.
(A) and (B) Schematic representations of the mutant tubg1 and tubg2 loci indicating the position of the T-DNA insertion sites. The structure of the three
loci was determined by DNA gel blot analysis, using T-DNA left and right border probes and sequencing of the T-DNA flanking regions (data not shown).
A single full-length T-DNA inserted in the first exon of the TUBG1 gene is present in tubg1-1 (A). A pair of T-DNAs organized in inverted repeat is inserted
in tubg2-1 (B). The T-DNA insertion induced a 2.4-kb deletion at the tubG2 loci, removing most of the coding region of TUBG2 from the end of the
second exon to the end of the gene 40 bp upstream of the transcriptional start of the adjacent At5g05630 gene. At5g05630 encodes an unknown protein
with weak similarity to amino acid permease family proteins. RT-PCR analysis showed that At5g05630 RNA is present in tubg2-1 plants at the same
level as in wild-type plants in all tissue examined (data not shown). This result, plus the fact that tubg2-1 homozygous plants have no visible phenotype,
strongly supports that the tubg2-1 insertion does not affect At5g05630 function. In the tubg2-2 line, three full-length linked T-DNAs are inserted in the
TUBG2 59 untranslated region. Gray boxes numbered I to X, exons; unnumbered gray boxes, 59 or 39 untranslated regions.
(C) RT-PCR analysis of TUBG1 and TUBG2 transcripts in wild-type organs and in mutant plants. Primer combinations specific for TUBG1 or TUBG2
cDNAs were used for RT-PCR amplification. The constitutively expressed APRT1 gene was used as a semiquantitative control (Moffatt et al., 1994).
Using a TUBG2-specific primer combination, an abundant fusion transcript is detected in the tubg2-2 mutant (bottom panel, arrow) that is also detected
using a T-DNA primer and a TUBG2-specific primer (right panel, arrow). Similarly, a fusion transcript between the T-DNA and the 39 part of the TUBG1
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combination induced severe gametophytic defects, and double
mutant seedlings were never recovered. For the other allelic
combination, tubg1-1 tubg2-2 double mutant seedlings displayed morphological defects and developmental arrests a few
days after germination. Detailed analysis of the phenotypes is
given below.
The severity of the phenotypes observed (gametophytic lethality in one case and seedling lethality in the other) suggested a
drastic depletion of g-tubulin levels in both double mutant combinations. Transcript levels are generally poor indicators of
molecular defects in insertional mutants. Therefore, we used an
anti-g-tubulin antibody to determine protein levels in double mutant plantlets by protein gel blot analysis. To get sufficient mutant
material, we used tubg1-1 tubg2-2 double mutant seedlings of
5 to 7 d, the earliest stage where morphological defects are clear
enough to unambiguously distinguish mutants from wild-type
plantlets. This experiment shows that, in comparison with the
strong signal observed in wild types of the same age, g-tubulin is
undetectable in such double mutant plantlets (Figure 1D).
Taken together, these results establish the following: (1) Genetically speaking, the observed phenotypes are strictly linked to
simultaneous disruption of both g-tubulin genes since they appear when insertion alleles are combined and only in this case. In
addition, a TUBG2 cDNA expressed under the control of the
promoter of the 35S gene of Cauliflower mosaic virus fully complements the phenotype of the tubg1-1 tubg2-2 mutant (Figure
5C), which further demonstrates the link between the mutations
and the observed phenotypes. (2) The absence of defect in single
mutants indicate high, if not complete, functional redundancy
between TUBG1 and TUBG2. (3) g-Tubulin is drastically reduced
and below detection level in tubg1-1 tubg2-2 seedlings. Given its
even more severe defects, the tubg1-1 tubg2-1 combination
presumably represents a null mutant, consistent with the molecular nature of the tubg1-1 (coding sequence insertion plus a
55-bp deletion) and tubg2-1 mutations (large deletion, as opposed to a 59 untranslated region insertion in tubg2-2).
We set out to characterize the developmental and cellular
alterations in this mutant material. The strong tubg1-1 tubg2-1
mutant combination was used to study mitotic defects induced
by g-tubulin depletion during both male and female gametogenesis. A GFP–microtubule binding domain (MBD) marker
was introduced into the tubg1-1 tubg2-2 background to have a
more dynamic view of microtubule arrays in g-tubulin–depleted
cells during postembryonic development.
g-Tubulin Depletion Affects Gamete Transmission
tubg1-1 tubg2-1 double heterozygote F1 plants (hereafter referred to as F1 plants) had a reduced seed set. Mature siliques of
these F1 plants contained ;24.0% desiccated ovules (484/2015),
presumably harboring a defective female gametophyte, a figure
highly significantly different from the <1% observed in the wildtype control (18/1847). This suggested abortion of most double
mutant female gametes since this frequency is very close to the
25% expected in case of complete female gametophytic lethality.
To determine the transmission efficiency of female and male
tubg1-1 tubg2-1 gametes, we used F1 plants in reciprocal
crosses with wild-type plants. The progeny of these crosses
were genotyped by PCR to score the transmission rate of double
mutant gametes. When F1 plants were used as female, only 2.1%
(5/244) of the progeny carried both tubg1-1 and tubg2-1 mutant
alleles, instead of 25% expected for full transmission. When F1
plants were used as a male donor, the double mutant tubg1-1
tubg2-1 gametes represented 9.4% of the gametes transmitted
to the progeny (26/276 plants). Therefore, transmission of double
mutant gametes was reduced by >90% on the female side and
by >60% on the male side, showing that the tubg1-1 tubg2-1
double mutation drastically alters formation and functioning of
both male and female haploid gametophytes.
In order to detail such defects, we then followed gametophytic
development in tubg1-1 tubg2-1 plants. As tubg1-1 and tubg2-1
mutations are in different genetic backgrounds (in ecotypes
Wassilewskija [Ws] and Columbia [Col], respectively), all the
following experiments were conducted on F1 hybrid plants
[TUBG1/tubg1-1; TUBG2/tubg2-1] to ensure a constant genetic
background and compared with Ws/Col F1 hybrid plants as wildtype controls. In such plants, 25% of produced gametes are expected to carry mutations in both genes.
Gametophytic Defects Induced by g-Tubulin Depletion
We used a procedure enabling visualization of the fine structure
of the embryo sac by confocal microscopy (Christensen et al.,
1997, 1998). Using this method, ovules exhibit autofluorescence
with nucleoli appearing extremely bright, whereas cytoplasm and
nucleoplasm fluoresce moderately, and vacuoles do not autofluoresce (Figure 2).
We examined >200 mature ovules of both wild-type hybrids
and double heterozygous F1 plants. In wild-type plants, we
observed <1% abnormal gametophytes, all arrested at the
Figure 1. (continued).
gene is detected in the tubg1-1 mutant. R, roots; RL, rosette leaves; S, stem; CL, cauline leaves; F, flowers; ª, suspension cultured cells; LG, light-grown
whole seedlings; DG, dark-grown whole seedlings; Gn, genomic DNA.
(D) Protein gel blot analysis of g-tubulin content in tubg1-1 tubg2-2 mutant plants. Proteins were extracted from whole seedlings aged 1 week. Total
protein extract (150 mg) from wild-type (left) and double mutant plants (right) was loaded on a two-well preparative SDS-PAGE gel. The gel was
transferred onto a membrane, and a 16-lane multislot blot system was used for hybridization, each lane corresponding to ;15 mg of total protein. The
bottom panel shows Coomassie blue staining of the membrane, revealing the ribulose-1,5-bis-phosphate carboxylase/oxygenase (Rubisco) large
subunit as a loading control. The top panel shows wild-type (lanes 1 to 3) and tubg1-1 tubg2-2 double mutant (lanes 4 and 5) extracts probed with a
polyclonal anti-g-tubulin antiserum raised against the full-length tobacco (Nicotiana tabacum) protein. The purified antibody was used at a dilution of
1:4000 (lanes 1 and 4) or 1:8000 (lanes 2, 3, and 5). Specificity was demonstrated by competition with 20 nM purified recombinant g-tubulin (lane 3).
g-Tubulin (;53 kD) migrates in the same region as the Rubisco large subunit, and a faint cross-hybridization of the g-tubulin antiserum to Rubisco is
visible on the protein gel blot just below the specific band, which is not competed out by addition of purified recombinant g-tubulin (lane 3).
g-Tubulin Function in Arabidopsis
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one-nucleate stage. In pistils of F1 plants, 33 ovules over 219
(15.0%) displayed abnormal gametophytes with reduced numbers of nuclei (Figures 2G to 2J) with respect to the wild type
(Figure 2F). Among these, embryo sacs with four (Figures 2H and
2I) or three nuclei (Figure 2J) were most frequent (;5% each).
Fewer mutant gametophytes contained six (Figure 2G), two, or
one nuclei. Nucleoli in these abnormal gametophytes were often
larger than in the wild type (compare Figure 2F with Figures 2G to
2J), suggesting a difference in ploidy level. Presence of large
nuclei in tubg1-1 tubg2-1 gametophytes was confirmed after
clearing of tissues and observation by differential interference
contrast microscopy (data not shown).
To determine the stage where gametophytic defects first
appear, we compared several stages of ovule development between wild-type and F1 plants, starting from the one-nucleate
stage to the mature gametophyte. Gametophyte development is
synchronous within one pistil (Christensen et al., 1997), which
enabled us to use wild-type gametophytes as internal calibration
for developmental stages. At early stages, all ovules of F1 plants
were of wild-type appearance. At the eight-nucleate stage, before nuclear migration, 36 abnormal gametophytes were observed from 228 ovules (15.8%) in F1 plants. As above for mature
gametophytes, defects included abnormal number, position, and
appearance of nuclei.
Therefore, among mutant tubg1-1 tubg2-1 gametophytes (25%
of total), 3/5 (15.8%) present detectable morphological defects;
the remaining ones, although undistinguishable from the wild
type at this stage, are mostly nonfunctional, since genetic analyses had previously showed that >90% of tubg1-1 tubg2-1
female gametes were defective.
The observation of male gametogenesis by differential interference contrast microscopy in anthers of F1 plants revealed that
meiosis and tetrad formation was not notably impaired by the
tubg1-1 tubg2-1 double mutation. During later stages, however,
the fraction of male gametophytes showing abnormalities was
significantly higher in F1 plants compared with the wild type
Figure 2. Phenotype of tubg1-1 tubg2-1 Female Gametophytes.
Confocal images of wild-type control ([A] to [F]) and tubg1-1 tubg2-1
gametophytes ([G] to [J]) prepared as described by Christensen et al.
(1997) (1998). The bright fluorescent spots are nucleoli and reflect the
position of nuclei. All ovules have the same orientation: the chalazal pole
is on the left, and the micropylar pole is on the right. ch, chalazal pole;
mp, micropylar pole; dm, degenerating spores; v, central vacuole; an,
antipodal nucleoli; pn, polar nucleoli; ccn, central cell nucleus; en, egg
nucleolus; sn, synergids nucleoli. These images are projections of
several 1-mm optical sections. Bars ¼ 20 mm.
(A) Wild-type uninucleate gametophyte: shortly after meiosis, three
spores degenerate, whereas the one located at the chalazal pole
expands, giving rise to the functional gametophyte.
(B) The functional spore undergoes mitosis and produces a two-nucleate
gametophyte visible here. Shortly afterwards, vacuoles coalesce into a
large vacuole separating the two nuclei.
(C) Four-nucleate gametophyte after second mitosis; the two pairs of
nuclei are separated by a large central vacuole.
(D) Eight-nucleate gametophyte generated by the third mitosis. The eight
nuclei are indicated by arrowheads.
(E) Nuclear migration in the eight-nucleate embryo sac: the two polar
nuclei (previously one at the chalazal pole and one at the micropylar pole)
have migrated to the micropylar half of the gametophyte where they will
ultimately fuse to give the central cell nucleus visible in (F).
(F) After the third mitosis, cellularization begins, giving rise to the mature
female gametophyte composed of the egg cell, which will give the
embryo upon fertilization, the central cell, which results from the polar
nuclei fusion that will give the endosperm after fertilization, and the two
synergids and three antipodal cells. The antipodal cells undergo cell
death upon complete maturation.
(G) to (J) Examples of abnormal gametophytes observed in mature
ovules of a [TUBG1/tubg1-1; TUBG2/tubg2-1] plant (six nuclei in [G], four
nuclei in [H] and [I], and three nuclei in [J]; arrowheads indicate position
of antipodal nuclei). In these gametophytes, nuclei present at the central
position are mostly of equal size (as judged from the size of the nucleoli),
instead of a central cell nucleus and a significantly smaller egg cell nucleus
in the wild type (compare with central cell nucleus and egg nucleolus in [F]).
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Table 1. Pollen Defects in Wild-Type and Double Heterozygous
Plants
Pollen Nuclear
Content
Number of
F1 Pollen
Observed
Percentage
Trinucleate
Binucleate 1
Binucleate 2
Uninucleate
2126
64
29
6
95.55%
2.88%
1.30%
0.27%
Number of
Wild-Type
Pollen
Observed
Percentage
574
6
98.97%
1.03%
Nuclear content was scored by DAPI staining of mature pollen of wildtype and F1 plants (double heterozygous for tubg1-1 and tubg2-1).
Binucleate 1 corresponds to pollen with one diffuse and one condensed
nucleus, whereas binucleate 2 contains two condensed nuclei. The
difference in abnormal pollen frequency is highly significant between the
wild-type and the F1 plants (P < 0.001).
(Table 1), as revealed by 49,6-diamidino-2-phenylindole (DAPI)
staining (Figure 3). The most frequent aberrant class with respect
to the wild type (Figure 3A) was binucleate pollen, with one nucleus resembling a vegetative nucleus and the other nucleus
more condensed and similar to a sperm nucleus (Figures 3B and
3C). Some pollen had two diffusely stained nuclei (Figures 3D
and 3E), and very few abnormal pollen had only one diffusely
stained nucleus (Figure 3F). As judged by Alexander staining,
there is no sign of pollen lethality in anthers of F1 plants (see
Supplemental Figure 1 online), and double mutant pollen grains
are viable. Moreover, in vitro germination (see Supplemental
Figure 1 online) did not reveal any notable difference between F1
and wild-type pollen. Given the transmission rate obtained in genetic analyses, this indicates that, as was the case for the female
side, a high proportion of double mutant pollen is morphologically normal and able to produce a pollen tube but not competent
for efficient fertilization.
We conclude that tubg1-1 tubg2-1 female gametophytes can
proceed normally through the very first stages of development
(first and second mitosis) and begin to show severe abnormalities and developmental arrests after the second mitosis. This
presumably reflects progressive consumption of the maternal
(sporophytic) g-tubulin stock. The timing of appearance of defects is variable from one embryo sac to another, which suggests
differences in initial amounts and/or in turnover of g-tubulin. The
same kind of mitotic defects are noted on the male side, although
less pronounced, which could be related to the smaller number
of divisions during pollen development and/or to a smaller dilution
factor of sporophytic proteins due to a smaller cellular volume.
Microtubule Defects during Mutant Pollen Development
Cells of g-tubulin mutants isolated in animal, fungus, and yeast
exhibit aberrant microtubule organization (Oakley et al., 1990;
Horio et al., 1991; Sobel and Snyder, 1995; Sunkel et al., 1995).
Defects in nuclear division observed during tubg1-1 tubg2-1 gametogenesis are consistent with abnormalities in microtubule
nucleation and organization. We consequently used a-tubulin
immunolabeling to compare microtubule organization during mutant and wild-type (Figure 4; see Supplemental Figure 2 online)
pollen development, from the first division of meiosis to trinucleate
pollen. Observation of pollen in anthers of double heterozygous F1
plants revealed that microtubule organization is undistinguishable
from the wild type up to the uninucleate pollen stage. Thereafter,
abnormal microtubular mitotic structures (24 out of 78 figures
observed) were detected (Figures 4C to 4H), which were never
seen in wild-type pollen (Figures 4A, 4B, and 4I; see Supplemental
Table 1 online). A common defect was abnormal spindles, which
appeared bent or collapsed and associated with unaligned chromosomes along the equatorial plane (compare Figures 4A and 4B
with Figures 4C to 4F; see Supplemental Table 1 online). Another
defect consisted of dense accumulations of microtubules at the
cell’s periphery in two-nucleate pollen just exiting division, suggesting that these structures correspond to collapsed phragmoplast (Figures 4G and 4H; see Supplemental Table 1 online).
Therefore, the tubg1-1 tubg2-1 mutation clearly affects both spindle and phragmoplast function during pollen development.
In addition, our observations strongly suggest a loss of asymmetry in the mutant. In wild-type pollen development, the first
mitotic division is strongly asymmetric and generates two morphologically distinct nuclei. During this division, the phragmoplast tends to encircle the highly condensed chromosomes of the
future sperm cell (Figure 4I; see Supplemental Figure 2J online).
In abnormal pollen with collapsed phragmoplasts, both daughter
Figure 3. Nuclear Phenotype of tubg1-1 tubg2-1 Mature Pollen.
DAPI staining of mature pollen grains. The mature male gametophyte of
Arabidopsis (or pollen grain) is a tricelled structure containing two sperm
cells (the gametes) enclosed in a vegetative cell. During pollen development, a sporogenous cell undergoes meiosis I and II and produces a
tetrad of spores. The first division of the spore is an asymmetric mitosis
that gives rise to a vegetative and a sperm cell, which is then internalized.
The sperm cell divides once more to produce two gametes. Upon
pollination, the vegetative cell sustains growth of the pollen tube in the
female tissue, enabling delivery of male gametes to the female gametophyte and double fertilization. Bars ¼ 10 mm.
(A) Typical trinucleate mature pollen grain from a wild-type hybrid control
plant. The nucleus of the vegetative cell stains less densely than the two
nuclei of the sperm cells that are highly fluorescent.
(B) to (F) Abnormal pollen grain in an anther of a [TUBG1/tubg1-1;
TUBG2/tubg2-1] plant. Most of the abnormal pollen contains only two
nuclei ([B] and [C]), one appearing dispersed and similar to the vegetative one and the other one appearing densely stained. Some pollen
contain two identical nuclei (E) and some only one nucleus (F).
g-Tubulin Function in Arabidopsis
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nuclei are equally stained (Figures 4G and 4H). Moreover, a centrally positioned phragmoplast separating two equally dense nuclei
are clearly visible in some mutant pollen grains (compare Figure
4I with 4J). Disruption of division asymmetry in pollen has already been shown in the gemini pollen1 mutant disrupted in the
Figure 5. Phenotype of the Aerial Parts of Wild-Type and tubg1-1
tubg2-2 Double Mutant Seedlings.
Figure 4. Microtubule Organization during tubg1-1 tubg2-1 Pollen
Development.
Overlay images of anti-a-tubulin immunolocalization (green) and DAPI
staining (blue).
(A) to (F) Spindle organization during the first mitosis of pollen development in anthers of wild-type hybrids ([A] and [B]) and [TUBG1/tubg1-1;
TUBG2/tubg2-1] F1 plants ([C] to [F]). Metaphase (A) and anaphase (B)
spindle in wild-type pollen. Spindle morphology is greatly affected in
tubg1-1 tubg2-1 mutant pollen, and spindles appear as bent ([C] to [F])
or even collapsed (F) structures.
(G) and (H) Abnormal microtubular structures in early two-nucleate
pollen in [TUBG1/tubg1-1; TUBG2/tubg2-1] anthers. DAPI staining and
nuclear morphology indicate that nuclear division has been achieved and
thus strongly suggest that collapsed microtubular structures correspond
to failed phragmoplasts.
(I) and (J) Loss of asymmetric division during mitosis I in pollen of
[TUBG1/tubg1-1; TUBG2/tubg2-1] plants (J) compared with a wild-type
plant (I). During mitosis I, phragmoplast is shifted to the side of the pollen
in the wild-type (I), whereas in a tubg1-1 tubg2-1 pollen, the phragmoplast is in the center of the cell.
Images are stack of 1-mm optical sections. Bar ¼ 8 mm.
(A) and (B) Phenotype of 2-week-old wild-type (A) and tubg1-1 tubg2-2
double mutant (B) plants. Mutant plants are similar to the wild type for the
first 3 to 4 days following germination and subsequently display several
morphological defects, including reduced expansion and deformation of
the cotyledons and defective leaf formation.
(C) The phenotype of a tubg1-1 tubg2-2 double mutant is fully rescued
by a 35S-TUBG2 construct. Shown here is a 3-week-old plant homozygous for the tubg1-1 and tubg2-2 mutations and heterozygous for the
complementing construct.
(D) and (E) Toluidine blue–stained longitudinal section in the shoot apical
meristem of 10-d-old wild-type (D) and tubg1-1 tubg2-2 double mutant
(E) plants. In the wild type, the shoot meristem displays small, densely
stained cells (D). The corresponding region in mutant plants contains a
reduced number of larger and less densely stained cells, indicating that
meristem cells initiated differentiation (E). Similarly, cells in the leaf
primordia (asterisk) undergo differentiation, and leaf formation is inhibited.
(F) to (H) Scanning electron micrographs of the shoot apical meristem
region.
(F) Emergence of leaves at the shoot apex of a 7-d-old wild-type
seedling.
(G) At the same stage, no leaves are initiated in the tubg1-1 tubg2-2
double mutant.
(H) Abnormal leaves are eventually initiated in the tubg1-1 tubg2-2
double mutant but undergo little subsequent growth, as shown in this
micrograph of the meristem of an 11-d-old double mutant seedling.
Bars ¼ 5 mm in (A) to (C) and 200 mm in (D) to (H).
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GEM1/MOR1 gene encoding a member of the MAP215 family of
microtubule-associated proteins (Park et al., 1998; Twell et al., 2002).
Sporophytic Lethality Induced by tubg1-1
tubg2-2 Mutations
In contrast with gametophytic defects of the tubg1-1 tubg2-1
combination, the tubg1-1 tubg2-2 mutant combination allows
normal gametophytic development and fertilization. Double homozygous mutant seeds develop and germinate normally, producing tubg1-1 tubg2-2 plantlets, which are similar to the wild
type up to 3 d after germination. At this stage, double mutant
plantlets start to display several morphological defects, eventually leading to lethality after 3 weeks of in vitro culture. The first
observable anomaly is a reduction of the cotyledons’ expansion,
which remain small and distorted (Figure 5B) with respect to the
wild type (Figure 5A). After 7 d of in vitro growth, it becomes
apparent that meristem activity is also impaired, as no more than
two leaf primordia are initiated at the shoot apex (compare
Figures 5D and 5F with 5E, 5G, and 5H). These primordia
undergo little growth and form small abnormally shaped leaves
(Figure 5H). Consistent with defective leaf formation, the shoot
apical meristem region is highly perturbed in the double mutant:
the characteristic dome-shaped organization of the wild type
(Figure 5D) is absent in tubg1-1 tubg2-2 double mutants (Figure
5E). Rather, the apex consists of a reduced number of large
cells, reminiscent of what occurs in mutants affected in the
maintenance of stem cell population (Barton and Poethig, 1993;
Laux et al., 1996; Moussian et al., 1998). This is an indication for
cell division arrest in the entire shoot apex and initiation of cell
differentiation.
Similarly, root growth is inhibited after 3 to 4 d of in vitro culture
(Figure 6). In the wild type, the root apex can be subdivided into
three zones: the distal division zone, where cells divide and
undergo radial expansion; the elongation zone, where considerable longitudinal cell expansion occurs; and the differentiation
zone, marked by the emergence of root hairs (Dolan et al., 1993;
Sugimoto et al., 2000). In 3- to 4-d-old tubg1-1 tubg2-2 mutants,
the overall root organization is conserved, but the sequence of
events is perturbed: Compared with cells in the wild-type root tip
(Figure 6A), cells start to elongate in the division zone of double
mutant seedlings (Figure 6B), resulting in a twofold to threefold
increase in cell length. Together with inhibition of root growth,
this indicates a premature arrest of cell division in the mutant root
meristem. Concomitantly, differentiating cells in the root apex
undergo considerable radial expansion, resulting in swelling of
the root apex (compare Figures 6C and 6E with 6A and 6D). No
more growth occurs following maximal swelling, and root morphology does not evolve past this point. Reflecting the growth
arrest in the root apex, the root hair differentiation zone eventually reaches the distal part of the root (Figure 6E).
Microtubule Misfunction in tubg1-1 tubg2-2 Cells
To monitor the effects of g-tubulin depletion on microtubule
organization, we studied microtubule organization in tubg1-1
tubg2-2 mutants using a reporter protein (GFP-MBD) (Marc
et al., 1998). tubg1-1 tubg2-2 mutants expressing the GFP-MBD
reporter were recovered in the progeny of crosses between
[tubg1-1/tubg1-1; tubg2-2/TUBG2] or [tubg1-1/TUBG1; tubg2-2/
tubg2-2] plants and a GFP-MBD–expressing line (Camilleri et al.,
2002). Double mutant progenies carrying the reporter gene were
selected and observed by confocal microscopy.
During the first 3 d after germination, mitotic arrays, such as
preprophase bands, spindles, and phragmoplasts, were properly assembled during mitosis as in the wild type (data not shown).
Similarly, cortical microtubule arrays in differentiating cells of roots
or hypocotyls were indistinguishable in appearance from the wild
type. Therefore, at this stage of development, where no signs of
morphological alteration were visible in double mutant plants, the
tubg1-1 tubg2-2 combination of alleles did not prevent formation
of typical microtubule structures.
Starting at 4 d after germination, microtubule defects become
visible in dividing and differentiating cells (Figure 7). The mitotic
activity is reduced by ;60% in double mutant root tips at 8 d
after germination (see Supplemental Table 2 online), and premature differentiation of cells in the division zone is noted (Figure
6), indicating a severe block of cell division in the root meristem.
Time-lapse observations of several dividing cells indicate that
duration of the cell cycle in mutant cells is longer and more
Figure 6. Root Growth and Morphology Defects in the tubg1-1 tubg2-2 Double Mutant.
Comparison of confocal optical median sections of roots of 4-d-old wild-type (A) and mutant (B) plants and 7-d-old mutant (C) plants. In the mutant, cell
expansion occurs in the root meristem (arrowhead in [B]), indicating premature arrest of cell division. The root apex thereafter undergoes radial swelling
(C), as also shown by scanning electron micrographs of wild-type (D) and double mutant (E) 11-d-old seedlings. Roots were stained with FM1-43 in (A)
to (C). Bars ¼ 50 mm in (A) to (C) and 200 mm in (D) and (E).
g-Tubulin Function in Arabidopsis
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Figure 7. Microtubule Organization in Wild-Type and tubg1-1 tubg2-2 Plants.
(A) to (D) Mitotic microtubule arrays in root meristem cells of wild-type plants 4 to 7 d after germination. Microtubules are first organized in a cortical
preprophase band encircling the nucleus (arrowheads in [A]). After nuclear envelope breakdown, the mitotic spindle forms (B). At the end of anaphase,
the phragmoplast, constituted of two sets of opposing microtubules, appears between the spindle poles (C), expands as a ring (D), and eventually
reaches the edge of the cells.
(E) to (H) Microtubules arrays in root meristem cells of 4- to 7-d-old tubg1-1 tubg2-2 seedlings. Fewer mitotic microtubule arrays were observed at
these stages in the mutant root meristem. Observed arrays include highly distorted or bent spindles ([E] and [F]), abnormal asymmetric phragmoplast
(G), or static condensed stacks of microtubules (H).
(I) to (L) Progressive disorganization of microtubule arrays after 3 to 4 d of postembryonic growth in tubg1-1 tubg2-2 mutants. Three days after
germination, cortical microtubules in elongating root cells are organized normally in parallel arrays, perpendicular to the cell main axis (I). Thereafter, in
the root elongation zone and early differentiation zone of 4- to 10-d-old mutant seedlings, progressive disruption of cortical microtubule arrays occurs,
accompanied by radial swelling ([J] to [L]). Microtubule defects are first characterized by a loss of transverse alignment ([J] and [K]). Subsequently,
cortical microtubules are depolymerized (L), as indicated by the presence of short microtubules (arrowhead) or patches of GFP fluorescence (inset) and
eventually by a diffuse cytoplasmic GFP staining.
Microtubule arrays are labeled by expression of the GFP-MBD fusion protein and imaged by confocal microscopy. (A) to (H) are single optical sections;
(I) to (L) are stacks of multiple images taken 1 mm ([I], [J], and [L]) or 2 mm (K) apart. Bars ¼ 10 mm in (A) to (H) and 50 mm in (I) to (L).
variable than in the wild type (Figure 8; see Supplemental Figures
3 and 4 online). In these cells, spindles (Figures 7E, 7F, and 8B to
8G) and to a lesser extent phragmoplasts (Figures 7G and 8H to
8J) are strongly perturbed compared with the wild type (Figures
7A to 7D). Condensed microtubule stacks (Figure 7H) are also
observable either at the center of the cell or at the cell periphery,
likely deriving from collapsed spindles or phragmoplasts. Cortical arrays appear to be less affected (Figure 8; see Supplemental
Figures 3 and 4 online).
In the elongation and differentiation zones, we observed
disorganization of interphase microtubule arrays (Figures 7I to
7L) and severe morphological defects, such as root swelling and
radial cell expansion (Figures 6C and 6E). Microtubule defects
are first characterized by a progressive loss of parallel orientation
of microtubules in some cells of the elongation zone (Figures 7I
and 7J). Subsequently, microtubule orientation is lost (Figure 7K),
and microtubule arrays are depolymerized, leaving a diffuse cytoplasmic GFP staining (Figure 7L). Hence, in addition to its role
in the formation of mitotic microtubule arrays, plant g-tubulin is
required for proper organization of the cortical network during
postembryonic development.
DISCUSSION
Overlapping Roles in Plant Development for the Two
Arabidopsis Genes Encoding g-Tubulin
g-Tubulin has been shown to be essential for microtubule nucleation at MTOC in all species where it has been analyzed. In
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The Plant Cell
Figure 8. Time-Lapse Observation of a Dividing Cell in the tubg1-1
tubg2-2 Double Mutant Root Tip.
(A) Preprophase band of normal appearance.
(B) to (E) Abnormal mitotic spindle.
(F) to (H) Abnormal asymmetric phragmoplast with two sets of opposing
microtubules of unequal width. The late disorganized phragmoplast is
finally rejected to the cell periphery (H).
(I) and (J) Exit of mitosis without karyokinesis (only one nucleus in [I]) and
without cytokinesis (absence of new membrane and cell wall deposition
in [J]).
The duration of the cell cycle (at least 144 min) is longer than in wild-type
root tip cells (<90 min in Azimzadeh et al., 2001). This is mostly due to the
persistence of the abnormal spindle (80 min from [B] to [E]) compared
with duration of a typical spindle phase in the wild type (20 min in
Azimzadeh et al., 2001). Microtubule arrays are revealed by the GFPMBD fusion protein and imaged by confocal microscopy. All images are
stacks of multiple images taken 0.6 mm apart. Bar ¼ 10 mm.
this article, we have investigated the consequences of g-tubulin
deficiency on plant development and microtubule organization in
acentrosomal Arabidopsis cells.
The three insertion lines used in this study all display normal
phenotypes in terms of gametogenesis, embryogenesis, and further seedling and plant development. However, combining mutations in both g-tubulin genes leads to severe developmental
defects and, depending on the allelic combination assayed, to
gametophytic or sporophytic lethality. This demonstrates that
the two Arabidopsis g-tubulin genes have overlapping, if not fully
redundant functions, as was predictable from their high sequence identity and overlapping expression patterns.
The molecular defects of the tubg1-1 and tubg2-1 alleles (i.e.,
deletion of almost the entire coding sequence for tubg2-1 and
an insertion plus a 55-bp deletion for tubg1-1) likely induce
complete loss of gene function in double mutant cells, consistent with the gametophytic lethality observed. Indeed, combination of these two alleles blocks nuclear division at various
stages during gametophyte development and results in mainly
abnormal, nonfunctional gametophytes. Genetic transmission
of double mutant gametes is very poor, especially for the female
ones. Nevertheless, a small proportion of double mutant gametophytes is functional and can proceed through fertilization.
These variations suggest that a pool of g-tubulin from parental
sporocytes is carried over into the gametophytes; depending on
the amount and/or stability of this parental stock, gametophytes
are able to sustain cell division and proceed into their development up to a point where g-tubulin reaches a critical level. This
critical concentration must be significantly lower than the normal
physiological concentration since the g-tubulin pool of the
meiocyte is sufficient to sustain a few rounds of cell division.
The fact that development of the female gametophyte involves
three successive mitoses and a larger cellular volume could
explain why embryo sac development is more severely affected
than the pollen.
The tubg1-1 tubg2-2 combination of alleles induces a weaker
phenotype since gametogenesis, embryogenesis, and early seedling development are normal in a double mutant background.
However, thereafter, perturbation of cell division and elongation
strongly affects seedling development and leads to seedling lethality after 3 weeks. Comparison of phenotypes induced by the
two different combinations suggests that tubg2-2 is a leaky allele
and likely allows synthesis of some residual g-tubulin, undetectable by protein gel blot analysis but sufficient to sustain embryogenesis and early seedling development. Synthesis of residual
g-tubulin in tubg1-1 tubg2-2 cells likely comes from fusion transcripts generated by the T-DNA insertion.
g-Tubulin Deficiency Induces Mitotic Defects
Our results reveal a range of abnormal mitotic microtubular arrays during tubg1-1 tubg2-1 male gametophyte development,
including abnormally shaped spindles and collapsed phragmoplasts. This is confirmed by the study of microtubule organization
in tubg1-1 tubg2-2 seedlings, where the same types of mitotic
defects are seen in meristematic cells. Altogether, these results
indicate that g-tubulin is necessary during mitosis for proper formation and/or function of the spindle and phragmoplast. Defective spindles are a common feature of g-tubulin mutants of
several species: S. pombe, S. cerevisiae, and Drosophila. In all
cases, spindles do assemble but are generally abnormal in shape
and lead to chromosome segregation defects (Horio et al., 1991;
Sobel and Snyder, 1995; Sunkel et al., 1995; Marschall et al.,
1996; Spang et al., 1996). In A. nidulans, g-tubulin depletion completely abolishes nucleation of spindle microtubules (Martin et al.,
1997). Whether Arabidopsis g-tubulin is required for proper functioning of mitotic arrays, is only involved in microtubule nucleation, or both remains to be determined.
g-Tubulin Function in Arabidopsis
Defective spindles likely result in unequal segregation of
chromosomes and formation of abnormally sized nuclei and presumably aneuploid cells, as observed in root tip cells (see
Supplemental Figure 3H online). Similarly, spindle misfunction
during mutant pollen development, as exemplified by the observation of distorted spindles (Figure 4), likely results in spermatic
nuclei with unbalanced chromosome stocks. Such gametic defects are not expected to impair pollen tube growth, which relies
on the activity of the vegetative nucleus. Indeed, our observations show that a high proportion of double mutant pollen is morphologically normal and able to produce a pollen tube. However,
aneuploid gametes, even if correctly delivered to the embryo sac,
are not likely to produce viable fertilization products, a hypothesis consistent with the poor transmission of double mutant
male gametes observed in our genetic analyses.
g-Tubulin is clearly required for initiation and proper functioning of mitotic spindles, and its depletion can result in complete
inhibition of nuclear division and polyploid cells. The abnormally
large nuclei observed in female tubg1-1 tubg2-1 gametophytes
(Figure 8) could correspond to such polyploid nuclei that went
through multiple cycles of replication without division. Inhibition
of nuclear division and increased ploidy have already been
reported for g-tubulin mutants in A. nidulans (Oakley et al.,
1990; Martin et al., 1997; Jung et al., 2001), S. pombe (Paluh et al.,
2000), and Drosophila (Sunkel et al., 1995) as well as in S. pombe
cells expressing mutant forms of human g-tubulin (Hendrickson
et al., 2001).
g-Tubulin has also been shown to play an essential role in
mitotic checkpoint (Hendrickson et al., 2001; Prigozhina et al.,
2004), and mutant forms of g-tubulin inhibit anaphase A and induce a delay in mitosis, with cells reentering interphase without
dividing (Prigozhina et al., 2004). The variation in cell cycle length
observed in Arabidopsis tubg1-1 tubg2-2 root tip dividing cells
may reflect a similar function for g-tubulin in plants (Figure 8; see
Supplemental Figures 2 and 3 online).
g-Tubulin Deficiency Affects the Organization
of Interphase Microtubules
Analysis of the tubg1-1 tubg2-2 combination of alleles indicates
that in Arabidopsis, g-tubulin is also required for proper organization of cortical interphase microtubules in elongating cells. This
finding is in agreement with a role of g-tubulin in cortical microtubule nucleation on extant microtubules, as evidenced by
Murata et al. (2005), but do not exclude additional effects on
polymerization, organization, or dynamics of microtubules. Indeed, in S. pombe and A. nidulans, g-tubulin is involved in the
spatial arrangement of cytoplasmic microtubules in addition to
its documented role in nucleation (Paluh et al., 2000; Jung et al.,
2001). Nucleation, dynamics, and spatial organization of microtubules are tightly coupled processes in plant cells and seem
all affected in tubg1-1 tubg2-2 mutants. In double mutant
cells, we observed fragmentation of cortical microtubules and
ultimately their complete disappearance, consistent with defective nucleation and alteration of dynamic properties of microtubules.
The phenotypic syndrome displayed by tubg1-1 tubg2-2 plantlets seems to be a common feature of plants with perturbed mi-
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crotubule arrays. For example, similar cytoskeletal defects and
root swelling are induced both by oryzalin and taxol, despite their
opposed effects on microtubule stability (Baskin et al., 1994), and
root swelling is also induced by the mor1 mutation (Whittington
et al., 2001) as well as in plants partially depleted in a-tubulin (Bao
et al., 2001). The precise timing of appearance of defects in the
tubg1-1 tubg2-2 weak combination at 3 to 4 d after germination is
intriguing. At this stage, residual g-tubulin synthesis either stops
or, in any case, becomes no longer sufficient to sustain growth and
development. A similar timing in the appearance of a root swelling
phenotype has been reported in Arabidopsis plants engineered for
reduced expression of a-tubulin genes (Bao et al., 2001). Bao et al.
(2001) proposed that the phenotype appears as roots attain a
maximal growth rate. The similarity with the mor1 phenotype
(Whittington et al., 2001) is in accordance with a role of g-tubulin in
the control of dynamic properties of cortical microtubules. The
mor1 temperature-sensitive mutation affects a plant homologue of
the TOGp/XMAP215 family of microtubule-associated proteins.
When shifted at restrictive temperature, cortical microtubules in
the mor1 mutant first lose their parallel alignment and eventually
depolymerize. This correlates with radial swelling of the root
(Whittington et al., 2001).
As in other eukaryotes, depletion of g-tubulin leads to severe
cellular defects and developmental arrests. These defects concerned all microtubule arrays, either mitotic, as exemplified by
defects observed during male and female gametogenesis, or interphase cortical arrays, as observed in a less severe, partially
viable mutant background. Minute amounts of g-tubulin seem sufficient for mitotic divisions since parental sporophytic stocks are sometimes sufficient to sustain formation of fully functional gametophytes.
Surprisingly enough, all Arabidopsis mutant cells observed
contained substantial amounts of microtubules, and defects in
such cells were rather related to organization and/or functioning
of microtubule arrays. In such mutant cells, where most probably
no de novo synthesis occurs, most residual g-tubulin is presumably titrated out of the cytoplasm and involved in g-TuRC complexes. Such nucleation complexes could allow de novo assembly
of microtubules to a certain extent, but the dynamic organization
of structured arrays may require constant spatial redistribution
and turnover of g-tubulin between the cytoplasmic pool and the
complexed form.
METHODS
Plant Material and Growth Conditions
The CVP11 (tubg1-1) and T628 (tubg2-2) lines derive from a T-DNA–
mutagenized population of the ecotype Ws obtained by the vacuum
infiltration procedure (Bechtold et al., 1993) using the Agrobacterium
tumefaciens strain MP5-1 carrying the transformation vector pGKB5
(Bouchez et al., 1993). The SALK_004612 (tubg2-1) line was obtained
from the ABRC and derives from a T-DNA–mutagenized population of the
Col ecotype (Alonso et al., 2003). GFP-MBD–expressing lines have been
described previously (Camilleri et al., 2002). Growth conditions are as
described by Nacry et al. (1998).
Molecular Techniques
Cloning, sequencing, and DNA gel blot analysis were performed essentially as previously described (Sambrook et al., 1989; Nacry et al., 1998).
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The Plant Cell
Plant DNA extraction was performed as described by Nacry et al. (1998).
For PCR screening of the Ws insertion lines, primers corresponding to
the T-DNA left border (TAG5: 59-CTACAAATTGCCTTTTCTTATCGA-39)
or right border (TAG3: 59-CTGATACCAGACGTTGCCCGCATAA-39) were
used in pair combinations with the following gene-specific primers: LeaderTubG1 (59-TCCTCACAGTCTCGAAACCC-39) and TubG2F0
(59-TACAAGTATTGTTAGAGAAG-39). Genotypes of the Ws mutant lines
were determined by PCR using the following primer pairs: LeaderTubG1/
TubGR01 (59-TATAGTGTTGGTCATCCG-39) for TUBG1; LeaderTubG1/
TAG3 for insertion in the tubg1-1 line; TubG2F0/TubGR01 for TUBG2;
and TubG2F0/TAG5 for insertion in the tubg2-2 line. Primer pairs used for
screening of the tubg2-1 line are as follows: TubG2F (59-CCTCTTCAGGCGTAGTAGTCTCGAAAC-39) and TubGR0 (59-TGTAGGGCTGGACAACAACGTCACT-39) for TUBG2 and TubG2F/LBb1 (59-GCGTGGACCGCTTGCTGCAACT-39) for the tubg2-1 line.
The probes used in DNA gel blot analysis of the tubg1-1 and tubg2-2
lines were a 3088-bp PstI-SstI fragment for the left border probe and a
2072-bp SstI-SstI fragment for the right border probe of the pGKB5
T-DNA (Bouchez et al., 1993). The probes used in DNA gel blot analysis
of the tubg2-1 insertion locus were a 457-bp PvuII-PvuII fragment (left
border probe) and a 1185-bp ApaI-NheI fragment (right border probe) of
the pROK2 T-DNA (http://signal.salk.edu/tdna_FAQs.html) and genomic
probes amplified from Arabidopsis thaliana genomic DNA with the
TUBG2F/TUBG2-7794 (59-GTCAATCATAACATTCAGAAGTCA-39) and
TUBG2-9280 (59-GAATGTGTTTTTTTTGGG-39)/TUBG2-17954 (59-TCTATAACGCCACTTAGC-39) primer pairs.
For RT-PCR analysis, single-stranded cDNAs were obtained from total
RNA as described by Camilleri et al. (2002). PCR was performed on
diluted cDNA samples using the primers LeaderTubG1/TubGRint1
(59-ACATCTTTTCTATCACCTCCCTGA-39) and TubG2F0/TubGRint1 for
amplification of TUBG1 and TUBG2 cDNAs, respectively. APT1 cDNA
was used as an internal control (Moffatt et al., 1994) and was amplified
using primers APT-RT1 (59-TCCCAGAATCGCTAAGATTGCC-39)/APTRT2 (59-CCTTTCCCTTAAGCTCTG-39).
Protein Extraction and Protein Gel Blot Analysis
To obtain antibodies directed against g-tubulin, a tobacco (Nicotiana
tabacum) cDNA was subcloned into pQE60 Escherichia coli His-tagged
expression vector and protein purified as recommended by the supplier
(Qiagen). Rabbit immunization was done according to Evrard et al. (2002)
using 200 mg of recombinant protein for each injection. IgGs directed
against g-tubulin were purified using thiophilic uniflow resin from Clontech. Purified IgGs were concentrated and stored at 208C in PBS containing 50% glycerol (v/v) and 1% BSA (w/v). Total protein extracts were
prepared as described by Liu et al. (1994). The protein concentration of
the extracts was determined using Bradford reagent (Sigma-Aldrich).
Proteins were separated on a 10% acrylamide gel and transferred to
Immobilon-P membranes according to the specification of the manufacturer (Millipore). The membranes were treated with rabbit anti-g-tubulin
antibody in the presence or absence of 20 nM recombinant g-tubulin. The
secondary antibody used was anti-rabbit IgG antibody linked to horseradish peroxidase (Sigma-Aldrich). Signals were revealed using the ECL
system (Amersham).
Anatomical and Indirect Immunofluorescence Microscopy
of Arabidopsis Gametophytes
Female gametophyte development was analyzed using a method developed by Christensen et al. (1997) (1998). The 488-nm laser line of an argon
laser of a Leica SP2 AOBS confocal laser scanning microscope was used
to illuminate and observe female gametophytes. Optical sections of 1 mm
were collected with a Leica HCX PL APO 363/1.20 NA water objective
and the Leica LCS software.
For the phenotypic analysis of pollen, four to five flowers were brushed
on a microscope slide. Pollen grains were observed under a Leica DMRB
microscope after Alexander staining (Alexander, 1969). DAPI staining
solution (100 mL) (0.1% Nonidet P-40, 10% DMSO, 50 mM PIPES, pH 6.9,
5 mM EGTA, pH 7.5, and 0.4 mg/mL of DAPI) was added to the slide, and
the pollen was covered with cover slips. The pollen was viewed by UV epiillumination using a Leica DMRB microscope. Pollen germination tests
were performed as described (Derksen et al., 2002).
Floral apical meristems were excised and were fixed, embedded, and
processed for immunofluorescence as described by Baluska et al. (2002).
Sections were incubated for 1 h with B-5-1-2 monoclonal anti-a-tubulin
(T5168; Sigma-Aldrich) diluted 1:400 (w/v) in PBS supplemented with 1%
BSA (w/v). After rinsing in PBS, sections were incubated for 1 h with the
secondary antibody Alexa Fluor 488 goat anti-mouse IgG (A-11017;
Molecular Probes) diluted 1:200 in PBS supplemented with 1% (w/v)
BSA. DAPI (1 mg/mL) was then applied for 10 min to label DNA. After
rinsing in PBS for 10 min, sections were mounted under cover slips and
were examined using a Leica SP2 AOBS confocal laser scanning microscope.
Anatomical Analysis of Arabidopsis Seedlings
Histological sections were prepared as follows. After overnight fixation in
4% paraformaldehyde in PBS, pH 7, samples were rinsed for 2 h in PBS
and dehydrated in a graded series of 10, 30, 50, 70, 90, and 100%
ethanol. Technovit 7100 (Kulzer, Heraeus) infiltration was performed according to the manufacturer’s instructions. Sections (5 or 10 mm) were
made on a Leica microtome and stained for a few seconds with 0.1%
toluidine blue. Sections were mounted in water and photographed on a
Leiz microscope. Seedling structure was studied using low-temperature
scanning electron microscopy as described by Traas et al. (1995). For
FM1-43 staining, living seedlings were stained with 2 mgmL1 FM1-43
(Molecular Probes) in water for 10 min and washed twice after staining,
and excised roots were mounted in water for observation. Imaging was
performed using a Leica TCS-NT confocal laser scanning microscope.
GFP-MBD fluorescence was observed on living seedlings mounted in
low-melting-point agarose (0.4% in water). Imaging was performed using
a Leica TCS-NT confocal laser scanning microscope.
Accession Numbers
Arabidopsis Genome Initiative locus identifiers for the genes mentioned in
this article are as follows: At3g61650 (TUBG1), At5g05620 (TUBG2), and
At1g27450 (APT1).
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Table 1. Mitotic Defects in Mutant Pollen.
Supplemental Table 2. Mitotic Defects in tubg1-1 tubg2-2 Roots.
Supplemental Figure 1. Pollen Viability from F1 Plants.
Supplemental Figure 2. Microtubule Organization during Male
Gametophyte Development.
Supplemental Figure 3. Time-Lapse Observations of Dividing Root
Tip Cells of the tubg1-1 tubg2-2 Double Mutant.
Supplemental Figure 4. Time-Lapse Observations of Dividing Cells in
the tubg1-1 tubg2-2 Double Mutant Root Tip.
ACKNOWLEDGMENTS
We thank the Salk Institute Genomic Analysis Laboratory for providing the sequence-indexed Arabidopsis T-DNA insertion mutants. We
g-Tubulin Function in Arabidopsis
also thank Daniel Vezon, Olivier Grandjean, and Christine Horlow for
technical advice, Richard Cyr for the gift of the GFP-MBD construct, and
Hervé Vaucheret for helpful discussions. J.A. was the recipient of a PhD
fellowship from the French Ministry for Research.
Received November 22, 2005; revised April 14, 2006; accepted April 21,
2006; published May 12, 2006.
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γ-Tubulin Is Essential for Microtubule Organization and Development in Arabidopsis
Martine Pastuglia, Juliette Azimzadeh, Magali Goussot, Christine Camilleri, Katia Belcram, Jean-Luc
Evrard, Anne-Catherine Schmit, Philippe Guerche and David Bouchez
Plant Cell; originally published online May 12, 2006;
DOI 10.1105/tpc.105.039644
This information is current as of June 15, 2017
Supplemental Data
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