Plant Cell Physiol. 44(3): 350–359 (2003)
JSPP © 2003
Randomization of Cortical Microtubules in Root Epidermal Cells Induces
Root Hair Initiation in Lettuce (Lactuca sativa L.) Seedlings
Hidenori Takahashi 1, Kayoko Hirota, Aiko Kawahara, Erika Hayakawa and Yasunori Inoue
Department of Applied Biological Sciences, Faculty of Science and Technology, Tokyo University of Science, Yamazaki 2641, Noda, Chiba,
278-8510 Japan
;
development by tip growth from a selected site on the bulge.
By analyzing morphological mutants of Arabidopsis, several
genes involved in the pathway leading to root hair formation
have been identified, including CPC (Wada et al. 1997), GL2
(Masucci et al. 1996), RHD2, 6 (Schiefelbein and Somerville
1990, Masucci and Schiefelbein 1994), and TTG (Galway et al.
1994). Cytological events during root hair formation have also
been well studied. It has been reported that the cell surface corresponding to the hair initiation site is acidified (Bibikova et al.
1998), and that a Ca2+ influx is important for directing hair tip
growth (Jones et al. 1995, Bibikova et al. 1997, Bibikova et al.
1999, Wymer et al. 1997). As with other intracellular events,
the cytoskeleton plays an important role in root hair formation.
Microtubules within the growing root hairs are distributed longitudinally or helically throughout the cytoplasm, and they
connect the subapical nuclei with the growing hair tips (Emons
1987, Lloyd et al. 1987, Ridge 1988). Bibikova et al. (1999)
revealed that microtubules regulate tip growth and orientation
in root hairs. Actin filaments have also been reported in root
hairs, and thick bundles of actin filaments are found over the
full length of the growing root hair (Traas et al. 1985, Lloyd et
al. 1987, Miller et al. 1999, Baluška et al. 2000, Tominaga et al.
2000). Actin filaments are thought to be involved in cytoplasmic streaming and the maintenance of transvacuolar strands
(Shimmen et al. 1995).
To understand the mechanism of root hair formation, both
root hair initiation (bulge formation) and root hair elongation
(tip growth from a bulge) must be studied. However, most cytological studies have focused on elongation. Little is known about
root hair initiation, because it is difficult to identify differentiating trichoblasts and hair-forming regions in the root. The subapical region, which is located just behind the zone of active
root elongation, is thought to be the hair-forming zone in most
plants (Esau 1965, Cutter 1978, Hofer 1996). However, some
reports suggest an overlap between the elongation zone and the
hair-forming zone (Cormack 1949, Jaunin and Hofer 1986).
In a previous study, we found that when lettuce seeds were
sown in liquid medium at pH 6.0 and pre-cultured for 24 h,
changing to fresh medium at pH 4.0 induced root hair formation, whereas changing to fresh medium at pH 6.0 did not
(Inoue et al. 2000). Root hair formation was initiated in a 1.5to 2.5-mm portion of the root tip by 4 h after acidification. The
hair-forming cells were located about 1.0 mm from the root tip
Root hair formation is induced when lettuce seedlings
are transferred from liquid medium at pH 6.0 to fresh
medium at pH 4.0. If seedlings are transferred to pH 6.0, no
root hairs are formed. We investigated the role of microtubules in this low pH-induced root hair initiation in lettuce.
At the hair-forming zone in root epidermal cells, microtubules were perpendicular to the longitudinal axis of the cell
just after pre-culture. This arrangement became disordered as early as 5 min after transfer to pH 4.0, and
became random by 30 min later. At pH 4.0, the randomization extended to the entire hair-forming zone of seedlings;
at pH 6.0, however, randomization did not occur and transverse microtubules were maintained. When seedlings at
pH 6.0 were treated with microtubule-depolymerizing
drugs, root hairs were formed. In contrast, when a microtubule-stabilizing drug, taxol, was added to the medium, no
root hairs formed, even at pH 4.0. These results suggest
that the transverse cortical microtubules inhibit root hair
formation, and that their destruction is necessary for initiation. Furthermore, the microfilament-disrupting drugs
cytochalasin B and latrunculin B inhibited root hair initiation, suggesting that actin filaments are necessary for root
hair initiation.
Keywords: Actin — Lactuca sativa L. — Low pH — Microtubule — Randomization — Root hair.
Introduction
Root hairs are tubular projections that develop in a specialized subset of root epidermal cells called trichoblasts. Root
hairs elongate by tip growth. They increase the surface area of
a root and hence its capacity to absorb water and nutrients.
Root hairs also play an important role in anchoring the plant to
the soil and in providing sites for interaction with a range of
symbiotic microorganisms (Clarkson 1985, Dolan et al. 1994,
Hofer 1996, Peterson and Farquhar 1996, Ridge 1996).
Root hair formation comprises at least two discrete
phases; the initiation phase, which is characterized by the
appearance of a small bulge in the outer periclinal cell wall,
and the elongation phase, which is characterized by root hair
1
Corresponding author: E-mail, [email protected]; Fax, +81-4-7123-9767.
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Microtubules during lettuce root hair initiation
351
Fig. 1 Organization of cortical microtubules in root epidermal cells. Seedlings at various stages were subjected to indirect immunofluorescence
microscopy using antibody against a-tubulin, and were observed using confocal laser scanning microscopy: just after pre-culture in medium at pH
6.0 for 24 h (A); after further culture with new medium at pH 4.0 for 15 min (B), 30 min (C) and 4 h (D, E), or at pH 6.0 for 4 h (F). The arrow in
D indicates the position of a bulge. Note that microtubules in the bulge are not observed in D since the focus is adjusted to the base of the bulge.
Scale bars = 10 mm.
at the end of the pre-culture (Inoue and Hirota 2000). Among
epidermal cells in the hair-forming zone, over 80% developed
root hairs by 7 h after acidification. These observations
revealed when and where bulge formation occurred in the main
root; as a result, changes in intracellular organization prior to
and during root hair initiation can be observed in a timedependent manner.
In this study, we examined lettuce seedlings to clarify the
role of the cytoskeleton, particularly the microtubules, in root
hair initiation. Indirect immunofluorescence microscopy was
performed using antibodies against tubulin, and the effects of
drugs that disrupt or stabilize microtubules were investigated.
Furthermore, we examined whether actin filaments are necessary for root hair initiation by applying inhibitors of actin filaments to the seedlings.
352
Microtubules during lettuce root hair initiation
Fig. 2 Effect of acidification on cortical microtubule organization in
root epidermal cells. Seedlings at various stages were subjected to
indirect immunofluorescence microscopy with antibody against atubulin: just after pre-culture in medium at pH 6.0 for 24 h (A); further culture with fresh medium at pH 4.0 for 5 min (B), 10 min (C),
30 min (D), and 4 h (E), or at pH 6.0 for 4 h (F). Angles between the
microtubule and the long axis of root cortical cells were measured for
over 15 seedlings and were classified into nine types (from 0° to 180°
at intervals of 20°). The number of microtubules with each range of
angles is indicated as a percentage.
Results
Microtubule organization during bulge formation
To clarify the role of microtubules in root hair initiation,
lettuce seedlings were subjected to indirect immunofluorescence
microscopy using antibody against tubulin. In our previous
study (Inoue and Hirota 2000), at the end of the pre-culture
hair-forming cells were limited to a region about 1.0 mm from
the root tip, and bulge formation began when the cells were
about 1.5 mm from the tip, 4 h after acidification. Therefore,
we examined microtubule organization in this hair-forming
zone. Just after pre-culture, cortical microtubules were perpendicular to the longitudinal cell axis in root epidermal cells (i.e.
the axis of the main root) (Fig. 1A). Fifteen min after the seed-
lings were placed in fresh pH 4.0 medium, the microtubules
became disordered (Fig. 1B). Thirty min after acidification, the
microtubules were completely randomized (Fig. 1C). The randomized microtubules were maintained until 4 h after acidification, when bulge formation began (Fig. 1D). The microtubules
were organized in mesh or strand shapes in the bulges (Fig.
1E). In contrast, when seedlings were placed in fresh medium
at pH 6.0, the transverse microtubules were maintained and
were observed even 4 h after changing the medium (Fig. 1F).
To analyze the transition of cortical microtubule arrays in
seedlings at pH 4.0 quantitatively, we measured the angle
between the cortical microtubule bundles and the longitudinal
axis of the root epidermal cells at different time points after
changing the medium. Just after pre-culture, 56% of the cortical microtubules in the epidermal cells of the hair-forming zone
were perpendicular to the long axis of the cell (Fig. 2A). The
number of transverse microtubules rapidly decreased to 39%
and 23% at 5 and 10 min after changing to pH 4.0 medium,
respectively (Fig. 2B, C). Thirty min after acidification, only
17% of the microtubules were transverse, and no difference
was observed in the numbers of microtubules at other angles
(Fig. 2D). These results indicate that microtubule arrays in the
root epidermal cells were completely randomized within
30 min of acidification. Randomized cortical microtubule
arrays were maintained until at least 4 h after acidification,
when bulge formation occurred (Fig. 2E). In contrast, when the
seedlings were placed in fresh medium at pH 6.0, no root hairs
formed, and the percentage of transverse microtubule arrays
was as high as 52%, even 4 h after changing the medium (Fig.
2F). The ratios of both transverse microtubules and microtubules at other angles were similar to the ratios observed just
after pre-culture (Fig. 2A). These results confirm that cortical
microtubules are randomized by acidification.
Relationship between cell position in the main root and cortical
microtubule randomization
After indirect immunofluorescence staining using antibodies against tubulin, root epidermal cells were classified into
five zones based on their distance from the root tip (0–0.9 mm,
0.9–1.2 mm, 1.2–1.5 mm, 1.5–1.8 mm, and >1.8 mm), and the
angles between the cortical microtubule bundles and the longitudinal cell axis were measured (Fig. 3). Just before changing
to pH 4.0 medium, microtubules in all epidermal cells were
most frequently observed to be transverse. More than half of
the cortical microtubules were arranged perpendicular to the
axis of cells at 0–1.8 mm from the root tip (Fig. 3A, C, E, G).
In cells more than 1.8 mm from the tip, transverse microtubules
were the most frequently observed but comprised only 30% of
the total (Fig. 3I). These microtubule arrays changed in a zonedependent manner after acidification. Four h later, the cortical
microtubule arrays were randomized in epidermal cells more
than 0.9 mm from the root tip (Fig. 3D, F, H, J). Epidermal root
cells located about 0.9–1.0 mm from the root tip are known to
sense low pH when medium is changed. After 4 h, they form
Microtubules during lettuce root hair initiation
353
Fig. 4 Cortical microtubules in root epidermal cells of seedlings
treated with colchicine. Seedlings were pre-cultured at pH 6.0 for 24 h
without colchicine and were transferred to fresh medium at pH 6.0
with 20 mM colchicine. They were harvested 4 h after the change of
medium and were subjected to indirect immunofluorescence microscopy with antibodies against a-tubulin. Epidermal cells without (A)
and with (B, C) root hairs are shown. A and B are fluorescent images.
C is a bright field image of B. Scale bar = 10 mm.
root hair bulges in a region 1.5–2.5 mm from the tip (Inoue and
Hirota 2000). Therefore, microtubule disruption was induced in
the entire hair-forming region. In contrast, cells located
between 0 and 0.9 mm from the root tip, which were covered
by the root cap and did not respond to the low pH, retained the
transverse cortical microtubule arrangement (Fig. 3B).
Fig. 3 Relationship between cell position in the main root and transition of the cortical microtubule organization in root epidermal cells.
Seedlings just after pre-culture with medium at pH 6.0 for 24 h (A, C,
E, G, I) and further culture with fresh medium at pH 4.0 for 4 h (B, D,
F, H, J) were subjected to indirect immunofluorescence microscopy
with antibodies against a-tubulin. Angles between the microtubule and
the long axis of root cortical cells were measured for more than 10
seedlings and summarized as in Fig. 2, depending on the distance from
the root tip (A and B, 0–0.9 mm; C and D, 0.9–1.2 mm; E and F, 1.2–
1.5 mm; G and H, 1.5–1.8 mm; I and J, >1.8 mm).
Effect of microtubule-disrupting and microtubule-stabilizing
drugs on root hair formation
The results described above indicated that cortical microtubule disruption was induced in root hair-forming epidermal
cells. However, it was unclear whether only microtubule randomization was necessary for root hair initiation, or whether
both acidification and randomization were necessary. If cortical microtubule randomization alone is enough to induce root
hair initiation, disruption of microtubules should cause root
hair formation in seedlings at pH 6.0. We therefore added
20 mM colchicine to the pH 6.0 medium after pre-culture and
observed its effects on root hair formation. Colchicine disrupted the microtubules completely (Fig. 4A), and many young
root hairs formed (Fig. 5A) that were indistinguishable from
those induced by pH 4.0 medium (Fig. 5I). Indirect immunoflu-
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Microtubules during lettuce root hair initiation
Fig. 5 Observation of the hair-forming zone in the main
root of seedlings with or without cytoskeletal drug treatment, using scanning electron microscopy. For colchicine, oryzalin, and propyzamide treatments, seedlings
were pre-cultured at pH 6.0 for 24 h without these drugs
and then were transferred to fresh medium at pH 6.0 with
20 mM colchicine (A), 10 mM oryzalin (B), or 10 mM
propyzamide (C). For taxol treatment, seedlings were
pre-cultured at pH 6.0 for 23 h without taxol, then cultured first at pH 6.0 for 1 h with 10 mM taxol and next at
pH 4.0 with 10 mM taxol (D). For cytochalasin B and
latrunculin B treatments, seedlings were pre-cultured at
pH 6.0 for 24 h without these drugs and were transferred
to fresh medium at pH 4.0 with 10 mM cytochalasin B
(E) or 1 mM latrunculin B (F). As controls, seedlings
were pre-cultured at pH 6.0 for 24 h and were transferred to fresh medium at pH 4.0 with 0.1% acetone (F)
or 0.1% DMSO (G), pH 4.0 without these solvents (E) or
pH 6.0 without these solvents (F). In each case, seedlings were harvested 8 h after the last change of medium
and were observed via low vacuum scanning electron
microscopy. Scale bar = 100 mm.
Microtubules during lettuce root hair initiation
355
Fig. 6 Effect of taxol on cortical microtubules of root epidermal
cells. Seedlings were pre-cultured at pH 6.0 for 23 h without taxol and
then cultured, first at pH 6.0 for 1 h with 10 mM taxol, and next at pH
4.0 with 10 mM taxol. Seedlings were harvested 4 h after the last
change of medium and were subjected to indirect immunofluorescence microscopy with antibodies against a-tubulin (A). Angles
between the microtubules and the long axis of root epidermal cells
were measured and summarized (B) as in Fig. 2. Scale bar = 10 mm.
Fig. 7 Effects of taxol and colchicine on main root growth. Seedlings were pre-cultured first at pH 6.0 for 23 h without taxol, then at
pH 6.0 for 1 h with 10 mM taxol, and finally at pH 4.0 with 10 mM
taxol (closed circle). Other seedlings were pre-cultured at pH 6.0 for
24 h without cytoskeletal drugs and were then placed in fresh medium
at pH 4.0 with 10 mM cytochalasin B (closed triangle), at pH 4.0 without cytoskeletal drugs (open circle), or at pH 6.0 without cytoskeletal
drugs (open triangle). Vertical axis indicates the increment of main
root length. Length at 0 h was designated 0 mm.
orescence staining confirmed that microtubules did not exist in
the colchicine-induced root hairs (Fig. 4B, C). Furthermore,
10 mM oryzalin and 10 mM propyzamide also disrupted microtubules (data not shown) and induced root hair initiation at
pH 6.0 (Fig. 5B, C). Since the oryzalin and propyzamide were
prepared with dimethyl sulfoxide (DMSO) as a solvent, 0.1%
DMSO was added to the pH 6.0 medium as a control. Root
hairs were not formed in DMSO-treated samples (data not
shown), suggesting that root hair formation was induced by
oryzalin and propyzamide and not by DMSO. These results
indicate that microtubules are not necessary for bulge formation and the early stage of root hair elongation. They also suggest that root hair initiation is induced only by the disruption of
transverse cortical microtubule arrays and that acidification is
not necessary for this process.
Using the microtubule-stabilizing drug taxol, we further
confirmed that the disruption of transverse cortical microtubules is essential for root hair initiation. If acidification (and
unidentified factor[s] such as positional information in the hairforming zone) is sufficient to induce root hair formation, root
hair induction should be observed in seedlings cultured at
pH 4.0 with taxol. When 10 mM taxol was added to pH 4.0
medium, transverse microtubules were still observed 4 h after
acidification (Fig. 6A); they comprised 51% of the total cortical microtubules (Fig. 6B). This percentage was in agreement
with that observed just after pre-culture (Fig. 2A). Furthermore, the percentages of microtubules at other angles were also
the same as those just after pre-culture (Fig. 2A), indicating
that cortical microtubule arrays were stabilized by taxol.
In contrast to the untreated seedlings, taxol-treated seedlings did not form root hairs even 8 h after treatment (Fig. 5D),
as in the case of untreated seedlings at pH 6.0 (Fig. 5J). Since
the taxol (and the cytochalasin B in the subsequent study) was
prepared with acetone as a solvent, 0.1% acetone was added to
the pH 4.0 medium as a control. Root hairs formed normally in
acetone-treated samples (Fig. 5G), suggesting that formation
was inhibited by taxol and not by acetone in the taxol-treated
seedlings. To examine the possibility that taxol inhibited root
hair formation by inhibiting main root growth rather than by
stabilizing transverse cortical microtubules, we measured the
speed of main root elongation. When seedlings were grown
with taxol at pH 4.0, the main root grew 2.0 mm in 8 h (Fig. 7,
closed circles). Slightly less growth (1.6 mm) was observed in
untreated seedlings at pH 4.0 (Fig. 7, open circles), whereas
more growth (4.0 mm) was observed in untreated seedlings at
pH 6.0 (Fig. 7, open triangles). These data indicate that root
356
Microtubules during lettuce root hair initiation
hair formation is inhibited not by main root growth inhibition
but by transverse cortical microtubule stabilization.
Effect of microfilament-disrupting drugs on root hair initiation
The microfilament-disrupting drugs cytochalasin B and
latrunculin B were used to investigate the role of actin filaments in root hair initiation. When seedlings were cultured
with 10 mM cytochalasin B at pH 4.0, root hair initiation was
inhibited and no bulges were observed in the hair-forming zone
(Fig. 5E). Root hair initiation was also inhibited by latrunculin
B (Fig. 5F). Since the latrunculin B was prepared with DMSO
as a solvent, 0.1% DMSO was added to the pH 4.0 medium as
a control. Root hairs formed normally in DMSO-treated samples (Fig. 5H), suggesting that formation was inhibited by
latrunculin B and not by DMSO in the latrunculin B-treated
seedlings. These results suggest that actin filaments are necessary for root hair initiation. In the presence of cytochalasin B at
pH 4.0, the main root grew 1.2 mm in 8 h (Fig. 7, closed
triangles), which is less than the growth seen in untreated
controls at pH 4.0 (1.6 mm; Fig. 7, open circles). However, we
previously reported that hair-forming cells are found 1.0 mm
from the root tip when the medium is changed, and that they
begin to form root hairs when they are 1.5 mm from the root tip
at pH 4.0 (Inoue and Hirota 2000). Since the elongation zone is
about 1.0 mm from the tip, growth of 1.2 mm is enough to
bring hair-forming cells to the 1.5 mm position. These observations indicate that root hair formation is inhibited not by main
root growth inhibition, but by inhibition of the polymerization
of actin.
Discussion
In this study, we found that randomization of transverse
cortical microtubule arrays occurred prior to bulge formation in
the hair-forming zone of lettuce seedlings at pH 4.0. In seedlings at pH 6.0, neither root hair formation nor cortical microtubule randomization was observed. However, at pH 6.0, seedlings formed root hairs when cortical microtubules were
disrupted. In pH 4.0 seedlings, transverse microtubule stabilization by taxol inhibited root hair formation. These results suggest that disturbance of the transverse microtubules by randomization or disruption is required to induce root hair
formation, and that acidification is not necessary. Therefore,
transverse microtubule arrays prevent root hair initiation.
Microtubules are known to determine the direction of cell
elongation by regulating the deposition of cellulose microfibrils. Since cellulose synthase moves along the cortical microtubules, the direction of cellulose microfibrils is parallel to the
cortical microtubules (Brown 1985, Delmer 1987, Herth 1985).
Cellulose microfibrils form a hoop, and cells expand perpendicular to the microfibrils. Therefore, the direction of cell elongation is perpendicular to the cortical microtubules (Lloyd
1982, Lloyd 1991, Gunning 1981, Roberts et al. 1985, Cyr
1994). The root epidermal cells of lettuce seedlings at pH 4.0
are shorter and more swollen than those of pH 6.0 seedlings,
suggesting that microtubules are involved in epidermal cell
elongation in the main root (Inoue and Hirota 2000). This interpretation is supported by the synchronization between the disarray of cortical microtubules (Fig. 3) and the reduction in the
elongation rate of the main root induced by acidification (Fig.
7, open circles, Inoue and Hirota 2000). When the microtubule
template is disrupted, cellulose deposition becomes disorganized and the direction of growth is lost (Baskin et al. 1994).
The shortening and swelling of epidermal cells at pH 4.0 may
be induced by the loss of directional elongation along the main
root axis caused by cortical microtubule randomization. The
slower elongation of the main root observed at pH 4.0 compared to pH 6.0 (Fig. 7) may reflect the shortening and swelling
of cells at pH 4.0.
Transverse cortical microtubules were also observed in
root epidermal cells of maize (Baluška et al. 1992) and Arabidopsis (Baluška et al. 2000). These microtubules may organize
the hoop of transverse cellulose microfibrils, which in turn
causes cells to elongate parallel to the main root axis and inhibits bulge protrusion from the lateral cell wall. In such situations, disorganization and relaxation of cellulose microfibrils is
required for bulge formation. Our observations of the induction of bulge formation by randomization or disruption of
transverse cortical microtubules are consistent with this
hypothesis. The importance of changes in cell wall organization during bulge formation is supported by recent studies.
Bernhardt and Tierney (2000) isolated At-PRP3, a gene encoding a proline-rich structural cell-wall protein that is expressed
during the later stages of root epidermal cell differentiation and
which is regulated by developmental pathways leading to root
hair outgrowth. Baumberger et al. (2001) reported that LRX1, a
chimeric leucine-rich repeat/extensin cell-wall protein, is also
required for root hair morphogenesis. In addition, a cellulose
synthase-like protein named KOJAK/AtCSLD3 is required at
the early stages of root hair outgrowth (Favery et al. 2001,
Wang et al. 2001). Cell-wall-modifying enzymes, including
expansins and xyloglucan endotransglycosylase, are also
known to be localized in outgrowing bulges (Baluška et al.
2000, Vissenberg et al. 2001).
Randomization of cortical microtubule arrays during
bulge formation was also observed in maize root (Baluška et al.
2000). In maize, trichoblasts are equipped with dense arrays of
transversely aligned cortical microtubules. These microtubules
are randomized and depleted as bulge outgrowth proceeds, and
randomization seems to occur only in the area of bulge formation. In lettuce seedlings, however, randomization is observed
in the entire hair-forming cell. Although the patterns differ in
these two species, these observations suggest that randomization of cortical microtubules is generally required for bulge formation, independent of plant species. Subjecting the maize root
to microtubule-stabilizing drugs in future studies will clarify
whether cortical microtubule randomization is essential for
bulge formation, as it is in lettuce seedlings. However, since
Microtubules during lettuce root hair initiation
only one bulge per cell formed at the appropriate position even
when all transverse cortical microtubules were randomized in
this study, microtubule randomization may be one of several
conditions necessary for bulge formation. Other factors may
include, for example, positional information in the cell.
Disruption of the cortical microtubule arrays produced the
same effect as randomization of the cortical microtubules by
acidification. Like the root hairs of acidified seedlings, lettuce
seedlings treated with colchicine, propyzamide, and oryzalin
formed straight root hairs, indicating that microtubules are not
required to regulate the directionality of tip growth or to maintain tip position in lettuce root hairs. However, microtubuledisrupting drugs cause waving or branching of root hairs in
Arabidopsis, cucumber, and tobacco, due to the loss of tip
growth directionality (Bibikova et al. 1999, Vissenberg et al.
2001). The involvement of microtubules in regulating the
directionality of root hair growth appears to vary from species
to species.
Recent studies have revealed that plant Rho-related
GTPases, termed Rop, localize to the future site of hair formation before bulge formation and to the tips of elongating root
hairs (Molendijk et al. 2001, Jones et al. 2002). The overexpression of Rop2 and constitutively active rop2 leads to additional and misplaced root hairs. Furthermore, dominant negative rop2 reduces the number of hair-forming sites (Jones et al.
2002). Conversely, Rop GTPase is thought to control the formation of cortical fine F-actin (Fu et al. 2002). These reports
suggest that Rop GTPase is involved in root hair initiation by
regulating actin dynamics. If so, actin filaments are needed for
bulge formation. In lettuce seedlings, actin filaments are
required for bulge formation, since cytochalasin B and latrunculin B inhibited root hair formation. This result supports the
importance of actin filaments in the process of root hair initiation. In Arabidopsis, Vicia, and maize, however, root hair initiation is thought to be actin independent, because bulges form
even when roots are treated with actin-depolymerizing drugs
(Miller et al. 1999, Baluška et al. 2000, Vissenberg et al. 2001).
In this respect, root hair initiation in lettuce seedlings is similar
to hair formation in lower plants, in which filamentous actin
patches or arrays mark the sites destined to initiate tip growth
(Bachewich and Heath 1998, Hable and Kropf 1998, Hable and
Kropf 2000, Alessa and Kropf 1999). Tominaga et al. (1997)
observed an elongated Hydrocharis root hair and proposed a
model in which microtubules are responsible for the organization of actin filaments in root hair cells. However, we revealed
that microtubules are not necessary for bulge formation,
although actin filaments are required. Our findings suggest that
the mechanism of organizing actin filaments differs in initiating
and elongated root hairs. Alternatively, actin filament alignment may be regulated differently in lettuce, and actin filaments may align in the appropriate direction by themselves
without microtubules.
Since the aim of this study was to investigate the role of
microtubules in root hair formation, we have not yet examined
357
how the formation of actin filaments changes during bulge formation. Future observation of actin filaments will provide
important insights into how cytoskeletons regulate root hair initiation.
Materials and Methods
Plant material
Virus-free lettuce (Lactuca sativa L. cv. Grand Rapids) seeds
were purchased from South Pacific Seeds (Griffith, N.S.W., Australia)
and kept in dry conditions at 4°C until use.
Culture conditions without cytoskeletal drugs
Seeds were immersed in tap water for 3 h under continuous white
light at 25°C to induce germination, and kept in the dark at 4°C for
24 h to synchronize the timing of germination. They were then sown
on a nylon mesh attached to a polystyrene frame and pre-cultured
hydroponically in 150 ml Arabidopsis medium (modified from Pruitt
[personal communication] as follows: 369.8 mg liter–1 MgSO4·7H2O,
472.2 mg liter–1 Ca(NO3)2·4H2O, 1,104.0 mg liter–1 NaH2PO4·2H2O,
303.3 mg liter–1 KNO3, 25.0 mg liter–1 EDTA-2Na, 2.4 mg liter–1
FeSO4·7H2O, 2.3 mg liter–1 MnSO4·4H2O, 0.24 mg liter–1 CuSO4·
5H2O, 0.29 mg liter–1 ZnSO4·7H2O, 1.86 mg liter–1 H3BO3, and
0.03 mg liter–1 (NH4)6Mo7O24·4H2O adjusted to pH 6.0 with NaOH).
This medium was poured into a polystyrene case and the culture was
kept under continuous white light at 25°C for 24 h to induce formation of the main root. Following the pre-culture, the old medium was
replaced by fresh medium at either pH 4.0 or pH 6.0 (adjusted with
HCl or NaOH, respectively), and the seedlings were then cultured for
different periods under continuous white light at 25°C. The maximum
pH change of the medium during 1-day culture was less than ±0.1.
Application of cytoskeletal drugs
Stock solution of colchicine (Wako Pure Chemical, Osaka,
Japan) was prepared at a concentration of 1,000´ in water. Stock solutions of taxol (Sigma, St. Louis, MO, U.S.A.) and cytochalasin B
(Sigma) were prepared at a concentration of 1,000´ in acetone. Stock
solutions of oryzalin (Wako Pure Chemical), propyzamide (Wako Pure
Chemical), and latrunculin B (Wako Pure Chemical) were prepared at
a concentration of 1,000´ in DMSO. For colchicine, propyzamide,
oryzalin, and cytochalasin B treatments, seeds were pre-cultured for
24 h in liquid medium at pH 6.0 without drugs (as described above).
The old medium was then replaced with fresh medium at pH 6.0 containing 20 mM colchicine, 10 mM propyzamide, or 10 mM oryzalin, or
fresh medium at pH 4.0 containing 10 mM cytochalasin B or 1 mM
latrunculin B. For taxol treatment, seedlings were pre-cultured for 23 h
at pH 6.0 without taxol and then cultured first at pH 6.0 with 10 mM
taxol for 1 h and then at pH 4.0 with 10 mM taxol.
Indirect immunofluorescence microscopy
Seedlings were fixed in freshly prepared 3.7% formaldehyde in
PIPES buffer [20 mM piperazine-1,4-bis(2-ethanesulfonic acid), 5 mM
EGTA, 5 mM MgCl2, 1% DMSO, 0.5 mM phenylmethylsulfonyl fluoride, pH 7.0] for 45 min and washed with PIPES buffer three times for
10 min each time. Seedlings were digested with 0.25% cellulase Onozuka R-10 (Yakult Honsha, Tokyo, Japan) and 0.025% pectolyase Y-23
(Yakult Honsha) in PIPES buffer and then washed with PIPES buffer
three times for 10 min each time. The digested samples were treated
with 1% Nonidet P-40 in PIPES buffer at 4°C for 5 h and washed with
phosphate-buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 4.3 mM
Na2HPO4, 1.4 mM KH2PO4, pH 7.2) three times for 5 min each time.
The samples were blocked with 2.5% bovine serum albumin (Sigma)
358
Microtubules during lettuce root hair initiation
in PBS at 25°C for 1 h. They were then incubated with mouse monoclonal antibody against a-tubulin (N356; Amersham Pharmacia Biotech, Amersham, Buckinghamshire, U.K.) at a dilution of 1 : 2,000 for
16 h at 4°C in blocking solution containing 0.25% sodium azide, and
washed in PBS three times for 5 min each time. The fluorescein-conjugated goat IgG fraction against mouse IgG (ICN Biomedicals, Aurora,
OH, U.S.A.) was diluted to 1 : 500 in blocking solution and then added
to the samples, which were incubated at 25°C for 1 h. They were then
washed in PBS three times for 5 min each and mounted on glass slides
in PBS supplemented with 0.1% p-phenylenediamine. The microtubules were observed by epifluorescence microscopy (E600; Nikon,
Tokyo, Japan) under blue-light excitation and with a MRC 600 confocal laser scanning microscope (Bio-Rad Laboratories, Hercules, CA,
U.S.A.) mounted on an Axiovert 135 (Carl Zeiss, Jena, Germany).
Scanning electron microscopy
Seedlings were harvested from the nylon mesh, and only the
roots were recovered. Surface water was removed with filter paper,
and the roots were fixed on an aluminum stage and immersed in liquid
nitrogen. The frozen roots were mounted on a cryo-stage holder and
observed with a low-vacuum scanning electron microscope (JSM5310LV, JEOL, Tokyo, Japan). Samples were kept at about –100°C
throughout. Images of the roots were observed using reflected electrons at about 30 Pa to visualize the outlines of rhizodermal cells.
Acknowledgments
The authors thank Dr. A. Kadota (Tokyo Metropolitan University, Tokyo, Japan) for his technical advice on indirect immunofluorescence microscopy. This work was supported by Grants-in-Aid to H.T.
from the Ministry of Education, Culture, Sports, Science, and Technology of Japan (Grant no. 13740474), the Foundation for the Advancement of Science Technology, and the Futaba Electronics Memorial
Foundation.
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(Received August 19, 2002; Accepted January 10, 2003)