Journal of Cell Science 104, 1217-1228 (1993)
Printed in Great Britain © The Company of Biologists Limited 1993
1217
A -tubulin-related protein associated with the microtubule arrays of
higher plants in a cell cycle-dependent manner
B. Liu1, J. Marc1,*, H. C. Joshi2 and B. A. Palevitz1,†
1Department
2Department
of Botany, University of Georgia, Athens, GA 30602, USA
of Anatomy and Cell Biology, Emory University School of Medicine, Atlanta, GA 30322, USA
*Present address: Biological Sciences A12, University of Sydney, Sydney, NSW 2006, Australia
†Author for reprint requests
SUMMARY
An antibody specific for a conserved -tubulin peptide
identifies a plant polypeptide of 58 kDa. -Tubulin antibody affinity purified from this polypeptide recognizes
the centrosome in mammalian cells. Using immunofluorescence microscopy, we determined the distribution of
this -tubulin-related polypeptide during the complex
changes in microtubule arrays that occur throughout
the plant cell cycle. We report a punctate association of
-tubulin-related polypeptide with the cortical microtubule array and the preprophase band. As cells enter
prophase, -tubulin-related polypeptide accumulates
around the nucleus and forms a polar cap from which
early spindle microtubules radiate. During metaphase
and anaphase, -tubulin-related polypeptide preferen-
tially associates with kinetochore fibers and eventually
accumulates at the poles. In telophase, localization
occurs over the phragmoplast. -Tubulin-related
polypeptide appears to be excluded from the plus ends
of microtubules at the metaphase plate and cell plate.
Its distribution during the cell cycle may be significant
in light of differences in the behavior and organization
of plant microtubules. The identification of -tubulinrelated polypeptide could help characterize microtubule
organizing centers in these organisms.
INTRODUCTION
Cande, 1990), for several fundamental reasons. First, the
acentriolar, anastral mitotic apparatus of plant somatic cells
is generally barrel-shaped, and spindle Mts at each pole do
not focus at a single discrete site but instead terminate at
separate loci containing membranes and diffuse amorphous
material (Hepler and Wolniak, 1984; Bajer and Molé-Bajer,
1986; Baskin and Cande, 1990; Lambert et al., 1991).
Second, interphase Mts do not emanate from a centrosome
close to the nucleus, as they do in animal cells. Instead,
they are largely restricted to the cortex in parallel arrays
that somehow control the orientation of new cellulose
microfibrils synthesized by plasma membrane-bound
glucan synthetases (Seagull, 1989; Shibaoka, 1991; Giddings and Staehelin, 1991). Third, at least two other arrays,
the preprophase band (PPB) and phragmoplast, appear at
different stages of the cell cycle and likewise are not linked
to a discrete, focused centrosome (Gunning and Wick,
1985; Seagull, 1989; Baskin and Cande, 1990; Wick, 1991).
It has been argued that the nuclear envelope (NE) in plants
serves as an MTOC for interphase, PPB and spindle Mts
(e.g. see Clayton et al., 1985; Wick, 1985; Seagull, 1989;
Flanders et al., 1990; Staiger and Lloyd, 1991), but definitive proof is still lacking (Hasezawa et al., 1991). In
“lower” plants, additional MTOCs are present at sites such
γ-Tubulin is a recently discovered member of the tubulin
superfamily (Oakley and Oakley, 1989) that appears to be
specifically localized to microtubule (Mt) organizing
(nucleating) centers (MTOCs) such as the centrosome of
animal cells (Stearns et al., 1991; Zheng et al., 1991) and
the spindle pole body of fungi (Oakley et al., 1990; Horio
et al., 1991). The protein, which shares approximately 30%
sequence homology with α and β-tubulins (Oakley and
Oakley, 1989), is present at relatively low abundance. Calculations indicate that all of the protein in an animal cell
may be restricted to the centrosome and none is incorporated into Mts per se (Stearns et al., 1991). Based on the
association with MTOCs, as well as genetic interactions
between γ-tubulin mutations and those of β-tubulin, it has
been proposed that γ-tubulin may nucleate and/or anchor
Mts and set up their characteristic minus (−) end proximal
polarity relative to the centrosome (Oakley et al., 1990).
In contrast to other eukaryotes, little is known about the
mechanisms governing the temporal and spatial control of
Mt formation in higher plants. Indeed, the nature of the centrosome in plants has long been debated (Wilson, 1925;
Mazia, 1984; Bajer and Molé-Bajer, 1986; Baskin and
Key words: centrosome, γ-tubulin, microtubule organizing
(nucleating) center, mitosis, phragmoplast, preprophase band
1218 B. Liu and others
as blepharoplasts and plastids (Hepler, 1976; Busby and
Gunning, 1989; Brown and Lemmon, 1990). To complicate
matters, proteins specific for MTOCs have not been identified in plants. While reports of cross-reactivity with antibodies directed against centrosomal moieties in other organisms have appeared (e.g. see Wick, 1985; Clayton et al.,
1985; Chevrier et al., 1992), the significance of these findings is either unclear or has been questioned (Harper et al.,
1989; Marc et al., 1989). Thus, the need for more information on MTOCs or centrosomal equivalents in plants is
imperative.
Joshi et al. (1992) have generated antibodies to a synthetic peptide corresponding to a conserved region in all
known γ-tubulins. This antibody recognizes γ-tubulin in cell
extracts, binds to the centrosome of mammalian cells,
blocks Mt growth from centrosomes in lysed cell preparations, and induces mitotic failure when injected into whole
cells. Because of the conserved nature of the peptide, we
reasoned that the antibody might cross-react with γ-tubulin
from plants. Our goal seemed reasonable, given that Stearns
et al. (1991) mentioned a partial γ-tubulin gene in maize.
We therefore used the antibody in immunoblot and
immunofluorescence studies and identified a γ-tubulinrelated polypeptide (GRP) in three plant species and two
cell types.
MATERIALS AND METHODS
-Tubulin antibody
All experiments were conducted with a purified immunoglobulin
(IgG) fraction (anti-γ) obtained from the polyclonal antiserum
(JH46) raised against the synthetic peptide EEFATEGTDRKDVFFYC coupled to keyhole limpet hemocyanin, as previously
described (Joshi et al., 1992). A similar IgG fraction obtained from
preimmune serum was used in controls.
Plant and animal material
Seedlings of Allium cepa L. cv. White Portugal (onion) were
grown in vermiculite, and Glycine max L. cv Williams 82 (soybean) in moist absorbent paper. Root tips were taken from 5- to
6- and 2-day-old seedlings, respectively. Suspension cultures of
Nicotiana tabacum L (BY-2) were maintained at 26°C according
to published procedures (see Hasezawa et al., 1991) and transferred weekly. Cells were harvested by centrifugation 2 days after
transfer. Gerbil fibroma cells (American Type Culture, line CCL
146) were obtained by courtesy of Dr. Charles Keith, University
of Georgia. The cells were grown on coverslips and culture dishes
in Dulbecco’s modified Eagle’s medium (Gibco-BRL, Grand
Island, NY) supplemented with 10% fetal bovine serum.
Preparation of material for immunofluorescence
observations
Root tips 1 mm in length were fixed for 1 h in freshly prepared
4% formaldehyde in PME (0.05 M Pipes buffer, pH 6.9, 5 mM
MgSO4 and 1 mM EGTA), rinsed several times in PME and partially digested for 30 min in a 1% Cellulysin (Calbiochem, San
Diego, CA) solution containing 5 mM EDTA, 20 µg/ml leupeptin,
5 µg/ml aprotinin, 5 µg/ml pepstatin A (all from Sigma Chemical Co., St. Louis, MO) and 50 µg/ml phenylmethylsulfonyl fluoride (PMSF) to retard protease activity. After further rinsing in
PME, root tip cells were released by gentle squashing onto slides
coated with gelatin and chrome-alum, affixed by air-drying and
then rehydrated in PME. Alternatively, cells were released onto
slides coated with polylysine (molecular weight 334,800; Sigma),
such that they remained wet throughout processing. Following
sequential treatment with 0.5% Triton X-100 for 30 min and
absolute methanol at −20°C for 10 min, the cells were treated with
antibodies.
Suspension cells were subjected to a somewhat different procedure. Following harvesting by centrifugation, the cells were
treated for 5 min with a 0.25 M mannitol solution containing 1%
Cellulase RS (Yakult Honsha Co. Ltd, Tokyo, Japan), 0.1% pectinase (Sigma) and the protease inhibitors listed above. After a brief
rinse in mannitol, the cells were allowed to adhere to polylysinecoated slides and then fixed in formaldehyde plus PME for 45
min. Rinsing in PME was followed by treatments with 0.5% Nonidet P-40 for 30 min, cold methanol for 10 min and rehydration
in PME.
Three days after plating, fibroma cells were fixed in cold
methanol for 5 min, rehydrated in PME and subjected to 0.5%
Triton for 10 min. Alternatively, the cells were prepared via the
formaldehyde procedure used for the plant cells.
Immunofluorescence staining
For anti-γ localizations alone, the antibody was diluted 1,600- or
3,200-fold (final protein concentration 2.5 or 1.25 µg/ml) in 3%
bovine serum albumin (Sigma) and 0.02% Tween-20 and applied
to slides or coverslips for 1 h or overnight at room temperature.
Following rinses in phosphate buffered saline (PBS), the cells
were treated for 45 min with FITC-conjugated goat anti-rabbit IgG
diluted 100-fold. Additional rinses in PBS were followed by
mounting in a medium containing 0.05 M Tris, pH 9.5, 1 µg/ml
Hoechst 33258 (Sigma) and 1 mg/ml p-phenylenediamine (Sigma;
an antifade agent).
For dual localizations, a mixture of 5 µg/ml monoclonal antiβ-tubulin (Boehringer-Mannheim, Indianapolis, IN) and 2.5 µg/ml
anti-γ was applied for 1 h or overnight. After rinsing in PBS, the
cells were treated with a mixture of Texas Red-conjugated sheep
anti-mouse IgG and FITC-conjugated goat anti-rabbit IgG, both
diluted 100-fold, for 45 min.
Immunofluorescence controls
In place of anti-γ, the IgG fraction of preimmune serum was
applied to cells at a 1,600-fold dilution. Alternatively, anti-γ was
boiled briefly before use. Finally, a taxol-stabilized brain tubulin
preparation (see below) was added to anti-γ at 20, 50 and 100
µg/ml concentrations before use on cells.
Microscopy and photography
Slides were examined on an Axioskop microscope (Carl Zeiss,
Thornwood, NY) equipped with epifluorescence optics. Images
were recorded on Tri-X film (Eastman Kodak, Rochester, NY)
and developed in HC-110 (Kodak).
Protein preparation
Allium and soybean root tips approximately 5 mm in length were
placed in a Dounce homogenizer and rinsed several times with distilled water. Washed roots were then covered with two volumes of
Laemmli sample buffer (Laemmli, 1970) containing 50-100 mM
dithiothreitol (DTT), 40 µg/ml PMSF, 2 µg/ml each of leupeptin,
chymostatin and pepstatin, and 10 µg/ml each of N-α-benzoyl-Larginine methyl ester (BAME; Sigma) and N-α-p-tosyl-L-arginine
methyl ester (TAME; Sigma) and homogenized with a teflon
plunger. Insoluble debris was sedimented at 12,000 g for 10 min
and the clarified protein extracts were heated at 85°C or boiled.
Gerbil fibroma cells grown in culture dishes were scraped off,
sedimented at 13,000 g for 30 s and solubilized with Laemmli
sample buffer as above.
γ-Tubulin in plants 1219
Bovine neuronal tubulin was prepared as described by Shelanski et al. (1973) and made MAP-free according to Cyr and Palevitz (1989). The tubulin was then polymerized with the aid of 10
µM taxol.
For reduction and alkylation, protein solutions were prepared
in Laemmli sample buffer containing protease inhibitors as
described above. Reduction with 50-100 mM DTT was carried
out at 85°C for 10 min. The samples were then alkylated at 85°C
for 5 min with iodoacetic acid dissolved in 0.5 M NaOH, pH 8.6,
according to Lane (1978) and Matsudaira (1989). The reaction
was quenched by adding β-mercaptoethanol to a 1% final concentration and the solution was adjusted to pH 6.8.
Gel electrophoresis and immunoblots
Proteins were separated on 8% acrylamide gels according to
Laemmli (1970), using an electrode buffer consisting of 24 mM
Tris, 192 mM glycine and 0.1% SDS. Separated proteins were
transferred electrophoretically onto nitrocellulose membrane (0.45
µm, BioRad Laboratories, Melville, NY) according to Towbin et
al. (1979) using a transfer buffer containing 50 mM Tris, 384 mM
glycine and 22% methanol. Transfer was carried out for 12-18
hours at 4°C and 200 mA. Nitrocellulose blots were incubated for
30 min with a blocking solution consisting of PBS, 1% BSA, 1%
lamb serum (Gibco-BRL) and 0.05% sodium azide.
Primary antibodies consisted of anti-γ diluted 500- to 1,000fold (in PBS containing 1% BSA, 10% lamb serum, 0.05% Tween
20 and 0.05% azide) and commercial monoclonal antibodies
against α and β-tubulin (Amersham Inc., Arlington Heights, IL)
diluted 1,000-fold. The anti-γ was followed by alkaline phosphatase-conjugated goat anti-rabbit IgG (Sigma) diluted 1,000- to
2,000-fold. The anti-α and β were first followed by rabbit serum
against mouse IgG (1 µg/ml) before treatment with the alkaline
phosphatase probe. The blots were developed with nitro blue tetrazolium and 5-bromo-4-chloro-3-indolyl phosphate (both Sigma)
substrate in carbonate buffer (100 mM NaHCO 3, 50 mM Na2CO3,
4 mM MgCl2, pH 9.5).
Affinity purification of anti-γ antibodies from the IgG fraction
was accomplished by using nitrocellulose blots as an affinity
matrix (Olmsted, 1981; Smith and Fisher, 1984). A large nitrocellulose blot was prepared from a gel loaded with a continuous
layer of fibroma proteins. Total proteins on the blot were visualized by staining with Ponceau S. The 49 kDa γ-tubulin polypeptide was identified by immunoblotting a narrow vertical strip from
the large blot with anti-γ, and the corresponding protein bands
were excised and destained. Blocking and antibody binding were
performed as before for immunoblots. Bound antibodies were
eluted with 0.2 M glycine at pH 2.9, neutralized with 1 M Tris,
pH 8.0, and dialyzed with PBS. As a control, the anti-γ IgG was
incubated with a nitrocellulose band taken from a non-reactive
protein on the large blot and eluted as above. Antibodies were
also affinity purified from the 58 kDa polypeptide in soybean
extracts.
RESULTS
Immunoreactive material detected by anti- shows
a punctate localization in microtubule arrays
Anti-γ produces distinct, consistent staining of Mt arrays in
all plant material examined (Fig. 1). While differences are
evident in the intensity of staining of each array between
sources, the patterns are very similar. Comparisons between
images produced by anti-γ and anti-β show that depending
on location or Mt array, the degree of coincidence with Mts
varies (see below). Moreover, anti-γ staining is finely punc-
tate (Fig. 1A,D). Although it is coincident with Mts, fluorescence is generally “rough” in appearance rather than the
continuous filamentous image produced with anti-β (compare Fig. 1A,D with B,E).
When we first applied anti-γ to root cells, we used the
dilutions (100- to 200-fold) reported for animal cells (Joshi
et al., 1992). However, the background signal was unacceptably high. We therefore diluted the antibody, whereupon fluorescence in association with Mt arrays became
obvious. Dilutions of 1,600- and 3,200-fold produced the
best contrast when applied to a variety of cells. The addition of BSA and Tween-20 to the diluent improved the
image as well, perhaps by reducing nonspecific binding.
However, fluorescent particles were always present to varying degrees, scattered throughout the cell regardless of the
staining protocol or cell type. Their significance, if any, is
unknown, but nonspecific particulate staining with another
γ-tubulin antibody has been noted previously (Oakley et al.,
1990).
Several other measures were taken to insure the specificity of staining. First, we used anti-γ on mammalian cells.
The resulting pattern was identical to that reported earlier
for animal cells using this antibody (Joshi et al., 1992), as
well as another anti-γ antibody (Stearns et al., 1992). Staining was primarily restricted to one or two dots in the region
corresponding to the centrosome, as ascertained from dual
localizations with anti-β (Fig. 2A,B). Single bright dots
were also present at the spindle poles of dividing cells (Fig.
2C,D). In addition, faint staining could be seen along the
proximal ends of spindle fibers radiating from the centrosome in metaphase and anaphase. In order to control for
the fact that the plant cells were fixed in formaldehyde and
the mammalian cells with cold methanol, we also subjected
the latter to formaldehyde. The staining pattern was identical: once again, localization over the centrosome was
prominent (data not shown), although the level of staining
was somewhat amplified.
Second, we performed several control localizations on
our plant cells. No staining except for a few bright particles scattered around the cell was seen with the IgG fraction of preimmune serum diluted to the same extent (Fig.
2E). Boiling anti-γ before use eliminated staining completely. On the other hand, inclusion of excess bovine brain
tubulin with anti-γ had no effect on staining (Fig. 2F).
Finally, we ascertained whether air-drying the plant cells
had any influence, by using material that was stuck to
polylysine-coated slides and therefore never dried before
staining. No difference was detected.
Anti- reacts with a 58 kDa GRP in plant material
In electrophoretic gels of protein extracts from onion and
soybean root tips, the α- and β-tubulins migrate to the
expected positions of about 52 and 54 kDa, respectively,
as shown on nitrocellulose blots incubated with monoclonal
antibodies (Fig. 3, lanes A,B). It is known that α and βtubulins run in reverse order on Laemmli gels compared
with their mammalian counterparts (see Fosket and Morejohn 1992). A polypeptide reacting with anti-γ is located in
the vicinity of the α- and β-tubulin subunits, at approximately 58 kDa (lane C). While this region seems to contain a doublet in the blot shown in Fig. 3, in other cases
1220 B. Liu and others
Fig. 1. Triple localizations of γ-tubulin (A,D), β-tubulin (B,E) and DNA (C,F) in Allium root cells. In the late anaphase cell shown in AC, anti-γ staining is seen near the spindle poles, in association with the clustered kinetochore fiber trunks (arrowheads in B). Little
staining is present along interzonal Mts. In the late telophase cell shown in D-E, only partial punctate anti-γ staining (D) is seen in
remaining portions of the phragmoplast (refer to E) next to the nearly completed cell plate. Note that the unstained midplane
encompassing the cell plate in D is wider than the cell plate proper in E. No specific pattern of anti-γ staining is evident elsewhere, despite
the presence of Mts around the daughter nuclei. Bar, 10 µm.
only a single band was seen. Thus, GRP runs behind the α
and β-polypeptides, in contrast to γ-tubulin in animal cells
(see below). No staining was seen with preimmune serum
(lane D).
Additional immunoreactive polypeptides appear at higher
molecular mass (e.g. 85, 100, 135 kDa) to varying degrees
in different samples. Such bands are present in boiled samples as well as those heated to 85°C. The polypeptides are
also present when samples were reduced and alkylated, and
when incubation of blots with the antibody was carried out
in the presence of 0.2% Triton X-100 and followed by
washing with a solution containing 0.5% Triton and 0.5%
SDS similar to that used previously on animal cells (Joshi
et al., 1992).
Similar to results obtained previously with other eukaryotic cells (Oakley et al., 1990; Stearns et al., 1991; Zheng
et al., 1991; Joshi et al., 1992), immunoblots of protein
extracts of mammalian cells detect a polypeptide at about
49 kDa, below the α and β-tubulin subunits at 56 and 54
kDa (Fig. 3, lanes F-H). The mammalian α, β and γ-tubulins thus migrate in descending order in standard Laemmli
gels. Furthermore, as in the immunoblots of plant proteins,
other polypeptides (e.g. at 100-108 kDa) are labeled.
Affinity-purified antibody detects centrosomes
and colocalizes with plant microtubules
In order to obtain mono-specific antibodies, we performed
small-scale affinity purifications of anti-γ using nitrocellulose blots of mammalian and plant proteins as an affinity
matrix (Olmsted, 1981; Smith and Fisher, 1984). The 49
kDa polypeptide of fibroma was identified by immunoblotting a narrow vertical strip from a large nitrocellulose blot.
Corresponding protein bands were excised from the large
blot, reacted with anti-γ, and bound antibody was eluted at
low pH. As a control, anti-γ was incubated with a nitrocellulose band from a non-reactive protein on the blot. The
affinity-purified antibody was then used for immunofluorescence microscopy as above.
As shown in Fig. 2G, antibody affinity purified from the
49 kDa polypeptide labels root cells in a manner similar to
that seen for the original anti-γ. Mammalian centrosomes
are also stained by this preparation (Fig. 2C). Interestingly,
a similar staining pattern was seen with antibody adsorbed
to one of the higher molecular mass polypeptides (100 kDa;
data not shown). No staining was seen with the control
eluate, however (data not shown). We also affinity purified
γ-Tubulin in plants 1221
Fig. 2. (A,B) Dual localization with anti-γ (A) and anti-β (B) in a
fibroma cell. A pair of bright dots is seen (A), corresponding to a
duplicated centrosome. The pair of dots is present at the
convergence point of cytoplasmic Mts (B). In (C and D) γ-tubulin
containing centrosomes are positioned at opposite ends of an
anaphase spindle, indicated by the Hoechst image of the
chromosomes in D. The anti-γ preparation used on this cell was
affinity purified from the 49 kDa polypeptide in fibroma extracts.
In (E) an Allium root cell treated with preimmune IgG is shown.
Only faint, nonspecific staining is seen. In (F) an Allium root cell
stained with anti-γ that had been pretreated with brain tubulin.
Staining is not diminished. In (G) an Allium root cell treated with
anti-γ affinity purified from the 49 kDa polypeptide of fibroma is
shown. The staining pattern in anaphase is identical to that seen
with non-affinity-purified antibody. In (H) centrosomes in a
fibroma cell are stained with anti-γ affinity purified from the 58
kDa polypeptide of soybean. The Hoechst image of the same cell
is seen in I. Bars, 10 µm.
Fig. 3. Immunoblots of protein extracts from soybean root tips
(lanes A-D) and fibroma cells (lanes E-H) probed with antibodies
against α (lanes A,F), β (lanes B,G) and γ-tubulin (lanes C,H) or
preimmune IgG (lanes D,E). The positions of molecular mass
markers (kDa) are indicated at the left and right. Note that A-D
and E-F were taken from different blots and gels.
are evident in the intensity of staining of each array between
plant sources, the overall patterns are very similar.
anti-γ from the 58 kDa polypeptide in soybean extracts. The
antibody still stains the centrosomes of mammalian cells
(Fig. 2H).
Cell cycle-dependent staining of plant Mt arrays
Immunoreactive staining is present in five distinct, cell
cycle-related locations. Fluorescence is associated with the
interphase cortical array, PPB, prophase NE, kinetochore
fibers and phragmoplast. Moreover, within each location,
the staining pattern changes over time. While differences
Interphase and prophase
Anti-γ positive material is concentrated in the cortex of cells
presumably in the G 1 phase of the cell cycle (Fig. 4A); only
a scattered, dispersed signal is seen elsewhere. This pattern
is most easily recognized in profile rather than face view.
Staining is clearly punctate in appearance, and in some
cases there is a hint that it occurs in lines or files. The intensity of the cortical signal varies, and in some cells it is
barely detectable (Fig. 4B). Cortical staining is quite evident in soybean (Fig. 4A), but less so in Allium. Comparison of coordinate anti-β images in dual-labeled material
1222 B. Liu and others
Fig. 4
shows that the weak anti-γ signal is associated with cortical Mts (Fig. 4B,C). This pattern is even more evident in
images obtained with Arabidopsis cells viewed on the confocal microscope (data not shown).
The PPB forms in the G2 phase of the cell cycle as an
γ-Tubulin in plants 1223
Fig. 4. (A-C) Distribution of anti-γ (A,B) and anti-β (C)
fluorescence in interphase root cells of soybean (A) and Allium
(B,C). A is from a single localization with anti-γ. A companion
Hoechst image (not provided) shows that the cell is in interphase.
Part of the cortex of another cell is also shown in A (*). B and C
are dual localization images at the same focal plane. Punctate antiγ fluorescence (arrowheads in A,B) is concentrated in the cortex.
The signal in A is much stronger than B, but comparison with
anti-β (C) indicates that it corresponds to cortical Mts. Inset in C
shows the Hoechst image of the interphase nucleus in B.
(D-F) Distribution of γ-tubulin (D), Mts (E) and chromatin in a
preprophase-prophase cell of soybean. Anti-γ fluorescence is
present in the preprophase band (arrowhead in D). Additional
staining is present around the nucleus, around which are numerous
Mts. (G-H) Triple localization with anti-γ (G), anti-β (H) and
Hoechst (I) in a preprophase-prophase Allium cell containing a
broad, split preprophase band, seen in face view. Anti-γ staining
of the lower segment of the band is prominent (arrowhead in G);
staining also is present but barely visible at the left side of the
upper segment. (J-K) Triple localization in a prophase soybean
cell. Anti-γ staining (J) is present in the narrowing PPB (K). Note
that fluorescence is also concentrating at the ends of the nuclear
envelope (*), a region of focused Mts. Bars, 10 µm (separate bars
for soybean and Allium).
initially wide band that gradually narrows as prophase
ensues (Palevitz, 1991; Wick, 1991). A concentration of
anti-γ staining is evident in early, wide PPBs (Fig.
4D,E,G,H). In addition, faint staining is also seen around
the nucleus (Fig. 4D). This perinuclear fluorescence is
notable because Mts accumulate at the nuclear periphery
around the time that the PPB appears, as is evident in Fig.
4E. As the PPB narrows, so does the area stained by antiγ (Figs 4J,K; 5A,B). In addition, the perinuclear fluorescence, which is initially uniformly distributed around the
nucleus, becomes progressively concentrated in cap-like
domains (Figs 4J; 5A,C). The caps correspond to the zones
upon which Mts begin to focus in prophase (Figs 4K;
5B,D). The caps are most striking in soybean, but less so
in BY-2 and Allium. Fig. 5E,F shows a broken BY-2 cell
in which anti-γ concentrates around a torus-like pole adjacent to the nucleus, while staining is less intense along Mts
extending away from this site.
The anti-γ signal in the PPB declines while Mts in the
band are still present (Fig. 5C,D). Nuclear cap fluorescence
remains prominent, however, and even increases. Eventually, PPB Mts disappear by prometaphase (Fig. 5H). As the
NE breaks down, anti-γ staining becomes more evident on
Mts emanating from the poles (Fig. 5G).
Metaphase-anaphase
As distinct kinetochore fibers are organized, fluorescence
seems more restricted to these elements (Fig. 6A-F). If
staining is present on Mts that branch from the kinetochore
fibers, it is very faint and difficult to detect against the dull
background fluorescence of the cytoplasm. Kinetochore
fiber fluorescence, like that seen earlier, is punctate. Interestingly, kinetochore fiber staining appears to be somewhat
restricted in that it does not extend all the way to the kinetochore. This is best appreciated when viewing most of the
kinetochore fibers simultaneously after dual localizations;
the nonstaining zone in the middle of the spindle encompassing the metaphase plate is wider in the anti-γ channel
Fig. 5. Dual anti-γ (A,C,E,G) and anti-β (B,D,F,H) localizations.
(A,B) Prophase soybean cell. Anti-γ staining is concentrated at
polar nuclear caps and in the preprophase band (arrowhead).
(C,D) Prophase soybean cell. Anti-γ staining is prominent in the
nuclear caps, but is now absent in the preprophase band.
(E,F) Staining of γ and β-tubulin at the torus-like nuclear poles (in
face view) in a BY-2 cell. (G,H) A prometaphase soybean cell.
Anti-γ staining is concentrated along Mts emanating from the
spindle poles. Inset in H: Hoechst image of the chromosomes.
Bars, 10 µm.
than in the anti-β channel (Fig. 6A-D). Staining of
metaphase spindles is weaker in Allium than soybean (Fig.
6E,F).
During anaphase, anti-γ fluorescence moves and shortens
1224 B. Liu and others
Telophase and the phragmoplast
Punctate staining is associated with the phragmoplast from
its inception in the interzone between daughter nuclei (Fig.
8A). Initially, as with the anti-β signal, the anti-γ positive
zone is broad, but then progressively narrows along with
the phragmoplast as it expands centrifugally (Fig.
8A,B,D,E). At the same time, staining decreases in older,
centripetal regions containing the cell plate (Fig. 8G,H).
The central plane occupied by the cell plate is unstained
(Fig. 8A,D). Again, the unstained central plane is wider
with anti-γ than anti-β, in a manner similar to that seen in
metaphase (compare Fig. 8A,B; D,E).
Weak anti-γ fluorescence is also present at the distal or
pole ends of the nuclei in Allium (Fig. 8D). Little or no fluorescence is seen associated with the Mts that emanate from
other regions of the NE at this stage (Fig. 8E). However,
more scattered perinuclear staining was noted in BY-2 (Fig.
8A).
DISCUSSION
Fig. 6. (A,B; C,D) Two metaphase soybean cells. Anti-γ staining
(A,C) is associated with the kinetochore fibers (B,D). Note that
the unstained region in the midzone in A and C is wider than the
metaphase plate seen with anti-β in B and D. (E,F) Dual
localization in a metaphase Allium cell. Punctate anti-γ staining
(E) is associated with the kinetochore fibers (F). Only weak
staining is seen along Mt branches. Bars, 10 µm.
with the kinetochore fibers (Fig. 7); it also becomes
brighter. The unstained region in the middle of the spindle
also widens, and in early anaphase, remains wider than the
midzone defined by the anti-β image (Fig. 7A,B). By midanaphase, the anti-γ fluorescence covers the entire kinetochore fiber and abuts the kinetochore, but is not present in
the interzone where anti-β shows extensive staining (Fig.
7C,D,E,F). Staining is progressively restricted to cap-like
aggregates at the poles in late anaphase, corresponding to
the stubs of kinetochore fibers (Fig. 1A,B). These caps are
especially prominent in Allium, but are clearly discernible
in soybean as well. Interzonal Mts stain only weakly if at
all (Figs 1A,B; 7). Eventually, staining at the poles becomes
fragmentary (Fig. 7G,H). Interestingly, Mts emanate from
the polar region at this stage (Fig. 7J; see Wick, 1985; Bajer
and Molé-Bajer, 1986).
Our results indicate that a peptide sequence unique to γtubulin is present in the cells of higher plants. Identification of a γ-tubulin-related protein is thus consistent with the
report of Stearns et al., (1991), who found a partial γ-tubulin gene sequence in maize. A γ-tubulin gene has also been
identified in Arabidopsis (C. Silflow and P. Snustad, University of Minnesota, personal communication).
The antibody used for this study reacts with γ-tubulin and
centrosomes in animal cells (Joshi et al., 1992). Our ability to confirm these results therefore allowed us more confidently to investigate GRP in plants. Further confidence in
our conclusions comes from the nearly identical results seen
in three different plant species and two cell types. Additional plant material has also been used, including Ara bidopsis cells, with very similar results. In addition, the
staining patterns in plant and mammalian cells were the
same after the antibody was cross affinity purified from the
fibroma and soybean polypeptides on nitrocellulose blots.
Staining at high dilution as well as the results of several
control experiments attest to the specificity of binding to
the plant cells. In particular, preadsorbtion with brain α and
β-tubulin does not suppress staining, indicating that the
antibody is not reacting with total tubulin.
On the other hand, there are clear differences in the distribution of GRP compared with that of γ-tubulin previously
reported in animal cells and fungi. First, the plant polypeptide migrates more slowly, at approximately 58 kDa, behind
α and β-tubulin. γ-Tubulin in the other species so far studied runs ahead of the α and β subunits. This could reflect
molecular mass differences, or it could represent an electrophoretic anomaly brought about by diversity in primary
sequence. It is also noteworthy in this regard that α-tubulin of plants runs ahead of β in Laemmli gels, as seen in
this study and documented earlier (Fosket and Morejohn,
1992).
While the γ-tubulin signal changes with the cell cycle in
animal cells, it remains largely restricted to the centrosome
(Stearns et al., 1991; Zheng et al., 1991, and the present
γ-Tubulin in plants 1225
Fig. 7. (A,B) Early anaphase soybean cell. Anti-γ fluorescence (A)
is concentrated around the kinetochore fiber trunks (B); only faint
staining is seen around interzonal Mts. Note that the unstained
interzone region is wider in A than in B. (C,D) Mid-anaphase
soybean cell. Anti-γ staining (A) is concentrated at the kinetochore
fibers trunks (B). The unstained interzone region is about the same
width in both views. (E,F) Mid-anaphase in an Allium cell. Anti-γ
fluorescence (E) is concentrated over the kinetochore fiber trunks
(F). Much fainter punctate staining is found associated with
interzonal Mts. (G) Anti-γ localization in a late anaphase cell in
Allium. Staining is concentrated at the poles. (H-J) Residual anti-γ
staining (H) at the poles in a late anaphase Allium cell. Little
fluorescence is seen elsewhere. A slightly different plane of focus in
J shows anti-β staining of Mts emanating from the poles. Hoechst
fluorescence of the chromosomes is shown in I. Bars, 10 µm.
results). Localization along Mts is faint and restricted to
portions of the spindle adjacent to the centrosome in mitotic
cells. Thus, the prominent association with Mts in plants is
at first surprising. It is noteworthy, however, that anti-γ does
not uniformly stain plant Mts. For example, Mts that branch
from kinetochore fibers and those in the spindle interzone
stain only weakly, if at all. The same can be said of those
that radiate along and/or from the NE at prophase and late
anaphase-telophase. Anti-γ staining of kinetochore fibers
stops short of the kinetochore proper, in contrast to anti-β.
Anti-γ also fails to stain the region immediately adjacent to
the cell plate, despite strong fluorescence with anti-β. Thus,
GRP seems to be excluded from the plus ends of Mts, an
observation which is consistent with the staining pattern in
animal cells. Interestingly, the kinetochores “catch up” to
the anti-γ staining in mid-anaphase, which is consistent with
an active role of the kinetochores in anaphase motion
(Gorbsky et al., 1987; Cassimeris et al., 1988).
The widespread distribution of anti-γ staining could represent the presence of the immunoreactive peptide in other
proteins; for example, in other tubulin subunits or in MAPs.
However, a computer search of data bases shows that the
peptide sequence is specific for γ-tubulin. Nevertheless,
higher molecular mass polypeptides are present to a variable extent in our plant samples. It is conceivable that the
bands represent γ-tubulin dimers. However, this is unlikely
given that the samples were run under denaturing conditions, the high molecular masses are not multiples of the
lower molecular masses, and the high molecular mass
polypeptides were seen in reduced and alkylated samples.
Thus, the significance of these polypeptides remains to be
ascertained.
The distribution of GRP in plants may have several
explanations. First, it is possible that the protein serves a
function different from that in animals. For example, it may
be incorporated as mixed dimers with α and β subunits into
Mt polymer. However, in that case a more uniform localization along Mts might have been expected, instead of the
punctate staining actually observed. On the other hand, the
distribution of GRP is consistent with a role in MTOC(s).
1226 B. Liu and others
Fig. 8. (A-C) An early phragmoplast in a BY-2
cell is shown in A and B. Punctate anti-γ
staining (A) is associated with the Mts (B).
Scattered additional fluorescence (arrowheads,
A) is seen around the re-forming nuclei
(Hoechst image, C). Note that the unstained
region bisecting the phragmoplast in A is wider
than the cell plate in B (arrowhead).
(D-F) Punctate anti-γ staining (D) in a late
Allium phragmoplast (E). Note that the
unstained mid-plane in D is narrower than in A,
but still wider than the cell plate in E.
Additional staining (*, in D) is seen at the distal
faces of the re-forming nuclei. The Hoechst
image of the nuclei is shown in F. (G,H) Face view of a late phragmoplast in soybean (G,H). Anti-γ fluorescence (G) is coincident with
the ring of phragmoplast Mts (H). Only dim anti-γ staining is seen in the middle of the cell, adjacent to older portions of the cell plate.
Bars, 10 µm.
Little is known about the spatial control of Mt organization
in plants, but there is reason to suspect that the ability to
nucleate Mts is dispersed in the cells of these organisms
(e.g. see Pickett-Heaps, 1974; Baskin and Cande, 1990).
First, no distinct centrosome is visible in somatic plant cells,
and the spindle poles are anastral (Bajer and Molé-Bajer,
1986; Baskin and Cande, 1990; Lambert et al., 1991). The
interphase Mt array is restricted to the cortex, and a variety of evidence indicates that it forms from dispersed cortical nucleation centers (e.g. see Cleary and Hardham, 1989;
Cho and Wick, 1989; Marc et al., 1989; Hasezawa et al.,
1991; Panteris et al., 1991). In addition, the NE (Seagull,
1989; Lambert et al., 1991) and phragmoplast (Zhang et al.,
1990; Vantard et al., 1990; Asada et al., 1991) also serve
to nucleate Mts at certain stages of the cell cycle. It can be
argued that the centrosome is a flexible entity that can
assume a variety of distributions in the cell (Mazia, 1984,
1987). Extension of this concept allows speculation that it
also dispersed during the evolution of sessile, terrestrial
plants in response to selection pressure in favor of the
development of a strong, complex cell wall containing
aligned cellulose microfibrils. Wall microfibrils are synthesized on the plasma membrane, and the mechanism controlling their orientation resides in the cortex in association
with Mts (Seagull, 1989; Giddings and Staehelin, 1991).
Given the need for an elaborate cortical cytoskeleton, it
seems logical that centrosomal elements necessary for its
formation should have dispersed as well, either at the NE
γ-Tubulin in plants 1227
or in the cortex itself. The evolution of the phragmoplast
(under the constraint of the wall as well as outwardly
directed turgor) perhaps also required additional MTOC
dispersal. Thus, the location of γ-tubulin became fragmented, and it could have become more generally associated with Mts as a result. It is noteworthy that additional
γ-tubulin-containing nucleation centers are found in fungi
as well (Horio et al., 1991).
An alternative hypothesis has been proposed in which
the behavior of plant Mts is intrinsically different than that
of animals in certain regards (Bajer and Molé-Bajer, 1986;
Smirnova and Bajer, 1992). Specifically, plant Mts commonly reorganize into aggregates or “converging centers”,
which can further develop into larger nucleating arrays. The
ability of plant Mts to form converging centers is ascribed
to several factors including specific properties of plant tubulin and/or minus ends (Bajer and Molé-Bajer, 1986;
Smirnova and Bajer, 1992). It is reasonable to suppose that
in the evolution of plants, γ-tubulin played a key role in
this behavior, thereby accounting for its more widespread
distribution. It is thus noteworthy that GRP still tends to be
excluded from the plus ends of Mts (e.g. in metaphase and
phragmoplast arrays), consistent with the pattern seen in
animal cells. Another phenomenon can also be explained
from the distribution of anti-γ staining: as the protein aggregates at the poles along with the shortening kinetochore
fibers, many additional Mts are generated from this region
in late anaphase (Wick, 1985; Bajer and Molé-Bajer, 1986).
The association of GRP with the NE appears to confirm
the importance of this structure in generating Mts. Considerable interest has been focused on the role of the NE in
this regard (Seagull, 1989; Lambert et al., 1991; Gunning,
1992). Indeed, anti-γ staining becomes prominent in
prophase when many Mts also appear around the nucleus.
Evidence favors the assumption that the minus ends of the
Mts are located at the NE (Vantard et al., 1990). Moreover,
the distribution of anti-γ around the NE changes in conformity with changes in the arrangement of the Mts and suspected MTOCs (Lambert et al., 1991). Specifically, it
becomes concentrated at the pole ends of the nucleus from
which Mts of the early spindle radiate (Baskin and Cande,
1990; Lambert et al., 1991). The staining pattern corresponds to the polar caps reported on prophase plant nuclei
by early cytologists such as Wilson (1925), who noted that
the caps give rise to the anastral spindle. However, while
Mts are generated at the re-forming nucleus in telophase
Allium cells (e.g. see Wick, 1985), perinuclear anti-γ staining is faint and declining. Thus, a better view of the function of this membrane awaits further experimentation.
Staining also appears in the midzone in late anaphasetelophase as the phragmoplast is organized, and its distribution changes as the phragmoplast expands toward the cell
periphery. While this distribution is consistent with the
phragmoplast operating as an MTOC, it is interesting that
γ-tubulin is not present immediately adjacent to the cell
plate, in the region of overlapping Mts. This distribution is
consistent with the plus ends of Mts being at the overlap
zone, with the minus ends away from it (Euteneuer et al.,
1982). This result is significant in light of recent counter
interpretations of tubulin assembly patterns in the phragmoplast (Vantard et al., 1990; Asada et al., 1991).
The presence of GRP in the PPB is of interest in light
of hypotheses concerning the origin of this band. Considerable evidence argues in favor of rearrangement of intact
Mts of the cortical array during band formation (e.g. see
Hasezawa et al., 1991; also see Palevitz, 1991; Wick, 1991).
It has also been proposed that the band arises via nucleation of new polymer at the band site in the cortex (e.g.
see Cho and Wick, 1989; Cleary and Hardham, 1989; Wick,
1991). A third possibility is that Mts arise at the NE (which
becomes active in Mt generation at this time) and are then
transported to the PPB site (Flanders et al., 1991) where
they are organized into parallel arrays. That GRP is present
in the PPB is significant, since it would favor the second
alternative (if indeed GRP is part of plant MTOCs). On the
other hand, the same pattern could arise if Mts are transported to the cortex and carry along with them part of the
MTOCs to which they were attached. Translocation and
reorganization of Mts (Bajer and Molé-Bajer, 1986; Palevitz, 1991) and accompanying nucleation factors could
explain much about changes in Mt distribution in plant cells
during the cell cycle.
Clearly, the distribution of GRP in plants changes in a
cell cycle-dependent manner, as does the distribution of Mts
and their organizing centers (Baskin and Cande, 1990; Lambert et al., 1991; Wick, 1991). Such changes may be attributable in part to alterations in GRP expression as well as
post-translational modifications. Investigations of GRP
mRNA levels therefore now are in order. Given the identification of a γ-tubulin gene from Arabidopsis (C. Silflow
and P. Snustad, personal communication), these experiments may come quickly. It will also be of interest to ascertain the number of γ-tubulin genes in plants, since, if more
than one is detected, they may be expressed differentially
in various Mt arrays. Preliminary experiments point to multiple γ-tubulin genes in maize (W.Z. Cande, personal communication). Experiments with two-dimensional PAGE
could shed further light on the occurrence of multiple GRP
isotypes, as they have with other tubulin families (e.g. see
Hussey et al., 1988).
We thank Dr. Richard J. Cyr, Pennsylvania State University,
for help with the preparation of brain tubulin, for useful discussions and for a computer search of protein sequences (with Dr
Mark Guiltinan); Dr Charles Keith, University of Georgia, for
kindly providing the fibroma cells; and Dr Seiichiro Hasezawa,
University of Tokyo, for supplying the original BY-2 cultures.
Soybeans were kindly provided by Ms Julie Lee and Dr Ronald
Nagao, University of Georgia. Ms Kimberly Duda helped print
micrographs. Supported by NSF grant DCB90-19285 to B.A.P.,
USDA grant 90-37261-5321 to J.M. and B.A.P., and NIH
(NS30009) and American Cancer Society (CD6255) grants to H.J.;
B.L. was also supported by a Research Assistantship from the
University of Georgia Graduate School.
REFERENCES
Asada, T., Sonobe, S. and Shibaoka, H. (1991). Microtubule translocation
in the cytokinetic apparatus of cultured tobacco cells. Nature 350, 238241.
Bajer, A. S. and Molé-Bajer, J. (1986). Reorganization of microtubules in
1228 B. Liu and others
endosperm cells and cell fragments of the higher plant Haemanthus in
vivo. J. Cell Biol. 102, 263-281.
Baskin, T. and Cande, W. Z. (1990). The structure and function of the
mitotic spindle in flowering plants. Annu. Rev. Plant Physiol. Plant Mol.
Biol. 41, 277-315.
Brown, R. C. and Lemmon, B. E. (1990). Monoplastidic cell division in
lower land plants. Amer. J. Bot. 77, 559-571.
Busby, C. H. and Gunning, B. E. S. (1989). Development of the
quadripolar meiotic apparatus in Funaria spore mother cells: analysis by
means of anti-microtubule drug treatments. J. Cell Sci. 93, 267-277.
Cassimeris, L. U., Walker, R. A., Pryor, N. K. and Salmon, E. D. (1988).
Dynamic instability of microtubules. BioEssays 7, 149-154.
Chevrier, V., Komesli, S., Schmit, A.-C., Vantard, M., Lambert, A.-M.
and Job, D. (1992). A monoclonal antibody, raised against mammalian
centrosomes and screened by recognition of plant microtubule organizing
centers, identifies a pericentriolar component in different cell types. J.
Cell Sci. 101, 823-835.
Cho, S.-O. and Wick, S. M. (1989). Micortubule orientation during
stomatal differentiation in grasses. J. Cell Sci. 92, 581-594.
Clayton, L., Black, C. M. and Lloyd, C. W. (1985). Microtubule
nucleating sites in higher plant cells identified by an auto-antibody against
pericentriolar material., J. Cell Biol. 101, 319-324.
Cleary, A. L. and Hardham, A. R. (1989). Microtubule organization
during development of stomatal complexes of Lolium rigidum.
Protoplasma 149, 67-81.
Cyr, R. J. and Palevitz, B. A. (1989). Microtubule-binding proteins from
carrot. I. Initial characterization and microtubule bundling. Planta 177,
245-260.
Euteneuer, U., Jackson, W. T. and McIntosh, J. R. (1982). Polarity of
spindle microtubules in Haemanthus endosperm. J. Cell Biol. 94, 644653.
Flanders, D. J., Rawlins, D. J., Shaw, P. J. and Lloyd, C. W. (1990).
Nucleus-associated microtubules help determine the division plane of
plant epidermal cells: avoidance of four-way junctions and the role of cell
geometry. J. Cell Biol. 110, 1111-1122.
Fosket, D. E. and Morejohn, L. C. (1992). Structural and functional
organization of tubulin. Annu. Rev. Plant Physiol. Plant Mol. Biol. 43,
201-240.
Giddings, T. H. and Staehelin, L. A. (1991). Microtubule-mediated
control of microfibril deposition: a re-examination of the hypothesis. In
The Cytoskeletal Basis of Plant Growth and Form (ed. C. W. Lloyd), pp.
85-99. London: Academic Press.
Gorbsky, G. J., Sammak, P. J. and Borisy, G. G. (1987). Chromosomes
move poleward in anaphase slong stationary microtubules that
coordinately disassemble from their kinetochore ends. J. Cell Biol. 104,
9-18.
Gunning, B. E. S. (1992). Use of confocal microscopy to examine
transitions between successive microtubule arrays in the plant cell
division cycle. Proc. Int. Symp. Cell. Basis Growth Dev. Plants (ed. H.
Shibaoka), pp. 45-55.
Gunning, B. E. S. and Wick, S. M. (1985). Preprophase bands,
phragmoplasts, and spatial control of cytokinesis. J. Cell Sci. 2, 157179.
Harper, J. D. I., Mitchison, J. M., Williamson, R. E. and John, P. C. L.
(1989). Does the autoimmune serum 5051 specifically recognize
microtubule organising centres in plant cells? Cell Biol. Int. Rep. 13, 471483.
Hasezawa, S., Marc, J. and Palevitz, B. A. (1991). Microtubule
reorganization during the cell cycle in synchronized BY-2 tobacco
suspensions. Cell Motil. Cytoskel. 18, 94-106.
Hepler, P. K. (1976). The blepharoplast of Marsilea: its de novo formation
and spindle association. J. Cell Sci. 21, 361-390.
Hepler, P. K. and Wolniak, S. M. (1984). Membranes in the mitotic
apparatus: their structure and function. Int. Rev. Cytol. 90, 169-238.
Horio, T., Uzawa, S., Jung, M. K., Oakley, B. R., Tanaka, K. and
Yanagida, M. (1991). The fission yeast γ-tubulin is essential for mitosis
and is localized at microtubule organizing centers. J. Cell Sci. 99, 693700.
Hussey, P. J., Lloyd, C. W. and Gull, K. (1988). Differential and
developmental expression of β-tubulins in a higher plant. J. Biol. Chem.
263, 5474-5479.
Joshi, H.C., Palacios, M. J., McNamara, L and Cleveland, D. W. (1992).
γ-Tubulin is a centrosomal protein required for cell cycle-dependent
microtubule nucleation. Nature 356, 80-83.
Laemmli, U. K.(1970). Cleavage of structural proteins during the assembly
of the head of bacteriophage T4. Nature 227, 680-685.
Lambert, A.-M., Vantard, M., Schmit, A.-C and Stoeckel, H. (1991).
Mitosis in plants. In The Cytoskeletal Basis of Plant Growth and Form
(ed. C. W. Lloyd), pp.199-208. London: Academic Press.
Lane, L. (1978). A simple method for stabilizing protein-sulfhydryl groups
during SDS-gel electrophoresis. Anal. Biochem. 86, 655-664.
Marc, J.M., Mineyuki, Y. and Palevitz, B. A. (1989). The generation and
consolidation of a radial array of cortical microtubules in developing
guard cells of Allium cepa L. Planta 179, 516-529.
Matsudaira, P. T. (1989). A Practical Guide to Protein and Peptide
Purification for Microsequencing. San Diego: Academic Press.
Mazia, D. (1984). Centrosomes and mitotic poles. Exp. Cell Res. 153, 1-15.
Mazia, D. (1987). The chromosome cycle and the centrosome cycle in the
mitotic cycle. Int. Rev. Cytol. 100, 49-92.
Oakley, C. E. and Oakley, B. R. (1989). The identification of γ-tubulin, a
new member of the tubulin superfamily encoded by mip A gene of
Aspergillus nidulans. Nature 338, 662-664.
Oakley, B. R., Oakley, C. E., Yoon, Y. and Jung, M. K. (1990). γ-Tubulin
is a component of the spindle pole body that is essential for microtubule
function in Aspergillus nidulans. Cell 61, 1289-1301.
Olmsted, J. B. (1981). Affinity purification of antibodies from diazotized
paper blots of heterogeneous samples. J. Biol. Chem. 256, 11955-11957.
Palevitz, B. A.(1991). Potential significance of microtubule rearrangement,
translocation and reutilization in plant cells. In The Cytoskeletal Basis of
Plant Growth and Form (ed. C.W. Lloyd), pp. 45-55. London: Academic
Press.
Panteris, E., Galatis, B. and Apostolakos, P. (1991). Patterns of cortical
and perinuclear microtubule organization in meristematic root cells of
Adiantumcapillusveneris. Protoplasma 165, 173-188.
Pickett-Heaps, J. D. (1974). Plant microtubules. In Dynamic Aspects of
Plant Ultrastructure (ed. A.W. Robards), pp. 219-255. London:
McGraw-Hill.
Seagull, R. W. (1989). The plant cytoskeleton. CRC Crit. Rev Plant Sci. 8,
131-167.
Shelanski, M., Gaskin, F. and Cantor, C. (1973). Microtubule assembly in
the absence of added nucleotides. Proc. Nat. Acad. Sci.USA 70, 765-768.
Shibaoka, H. (1991). Microtubules and the regulation of cell
morphogenesis by plant hormones. In The Cytoskeletal Basis of Plant
Growth and Form (ed. C. W. Lloyd), pp. 159-168. London: Academic
Press.
Smirnova, E. A. and Bajer, A. S. (1992). Spindle poles in higher plant
mitosis. Cell Motil. Cytoskel. 23, 1-7.
Smith, D. E. and Fisher, P. A. (1984). Identification, developmental
regulation, and response to heat shock of two antigenically related forms
of a major nuclear envelope protein in Drosophila embryos: application
of an improved method for affinity purification of antibodies using
polypeptides immobilized on nitrocellulose blots. J. Cell Biol. 99, 20-28.
Staiger, C. J. and Lloyd, C. W. (1991). The plant cytoskeleton. Curr. Opin.
Cell Biol. 3, 33-42.
Stearns, T., Evans, L. and Kirschner, M. (1991). γ-Tubulin is a highly
conserved component of the centrosome. Cell 65, 825-836.
Towbin, H., Staehelin, T. and Gordon, J. (1979). Electrophoretic transfer
of proteins from polyacrylamide gels to nitrocellulose sheets: procedure
and some applications. Proc. Nat. Acad. Sci. USA 76, 4350-4354.
Vantard, M., Levilliers, N., Hill, A.-M., Adoutte, A. and Lambert, A. M.
(1990). Incorporation of Paramecium axonemal tubulin into higher plant
cells reveals functional sites of microtubule assembly. Proc. Nat. Acad.
Sci.USA 87, 8825-8829.
Wick, S. M. (1985). Immunofluorescence microscopy of tubulin and
microtubule arrays in plant cells. III. Transition between
mitotic/cytokinetic and interphase microtubule arrays. Cell Biol. Int. Rep.
9, 357-371.
Wick, S. M. (1991). The preprophase band. In The Cytoskeletal Basis of
Plant Growth and Form (ed. C.W. Lloyd). London: Academic Press.
Wilson, E. B. (1925). The Cell in Development and Heredity, pp.12-32.
New York: MacMillan.
Zhang, D., Wadsworth, P. and Hepler, P. K. (1990). Microtubule
dynamics in living dividing plant cells: confocal imaging of
microinjected fluorescent brain tubulin. Proc. Nat. Acad. Sci. USA 87,
8820-8824.
γ-Tubulin in plants 1229
Zheng, Y., Jung, M. K. and Oakley, B. R. (1991). γ-Tubulin is present in
Drosophila melanogaster and Homo sapiens and is associated with the
centrosome. Cell 65, 817-823.
(Received 2 November 1992 - Accepted 11 December 1992)