Download C-Tubulin in Barley and Tobacco: Sequence Relationship and RNA

Plant Cell Physiol. 43(2): 224–229 (2002)
JSPP © 2002
C-Tubulin in Barley and Tobacco: Sequence Relationship and RNA Expression Patterns in Developing Leaves during Mitosis and Post-Mitotic Growth
Jan Schröder, Kordula Kautz and Wolfgang Wernicke 1
Institut für Allgemeine Botanik, Johannes Gutenberg – Universität Mainz, P.O. Box 3980, D-55099 Mainz, Germany
;
C-Tubulin is typically associated with microtubule
organising centres, such as the centrosome, and appears to
mediate microtubule nucleation. Centrosomes are usually
not found in higher plants, but active genes homologous to
C-tubulin have been identified in the plant kingdom, including the angiosperms Arabidopsis, maize and rice. We have
isolated and characterised C-tubulin cDNA sequences of two
further angiosperm species, barley and tobacco. Sequence
comparison revealed a phylogenetic tree with distinct clusters corresponding to the systematic position of the species.
Furthermore, domains, thought to be exposed in the folded
protein and to be candidates for interaction with associated,
nucleation-site related proteins, exhibited motifs highly specific of multicellular plants. Strong expression of the Ctubulin genes, as determined by Northern blotting, correlated with mitotic activity. Expression dropped distinctly
when mitotic activity ceased. Thus, in post-mitotic tissues
that showed intricate reshuffling of cortical microtubule
arrays related to cell shaping only very low C-tubulin
steady-state RNA levels were found, contrasting with the
situation for =-tubulin. The findings indicate that C-tubulin
expression in plants may be more tightly linked to mitosis,
although there is some C-tubulin expression at the RNA
level even after mitosis. It follows that the post-mitotic
changes in microtubular arrays may be less dependent on
concurrent C-tubulin RNA expression than mitotic cells.
ing in interphase and terminally differentiating post-mitotic
cells. Little is known as to how the successive establishment of
the different arrays is accomplished (Vaughn and Harper 1998).
There is increasing evidence that plant microtubules are prone
to rapid turnover, with interphase microtubules turning over
more than three times faster compared to those in animal cells
(e.g. Hush et al. 1994, Moore et al. 1997). This indicates that
assembly and disassembly cycles play a dominant role in rearrangement of microtubule arrays in plants, rather than shifting
of whole stable microtubules.
De novo assembly of microtubules in vivo appears to rely
on specialised nucleation sites, typically in the form of a
microtubule organising centre (MTOC). The dominant MTOC
is the centrosome of mammalian cells. Here, nucleation is most
likely mediated by g-tubulin, a member of the tubulin superfamily that is capable of forming complexes acting as templates
(Oakley and Oakley 1989, Moritz et al. 2000, Oakley 2000).
Such centrosome-like MTOCs are usually not observed in
higher plants, especially not in angiosperms (Vaughn and
Harper 1998), but genes homologous to g-tubulin have recently
been found in the plant kingdom, including some angiosperm
species, Arabidopsis (Liu et al. 1994), maize (Lopez et al. 1995)
and rice (Kim et al. 1999). Several attempts have been made to
detect the protein in cells by means of immunofluorescence
microscopy. It emerges that higher plant g-tubulin co-localizes
with most if not all of the microtubule patterns described above,
more or less mirroring the arrays, albeit in a punctate fashion
(e.g. Liu et al. 1993, Canaday et al. 2000, Panteris et al. 2000).
The findings leave open the question as to whether g-tubulin
acts in higher plants as a highly dispersed nucleation site or
whether it has evolved different functions (Vaughn and Harper
1998, Panteris et al. 2000). In this context it is important to note
that non-centrosomal microtubule nucleation has been observed
in animals (Hyman and Karsenti 1998, Oakley 2000).
We are interested in changes of microtubular arrays occurring during transition from cell division to cell differentiation.
The dynamics of microtubules in cycling cells are well established and indirect evidence suggests that even post-mitotic
changes in cortical microtubular arrays are not simply the
result of shifting whole stable microtubules (e.g. Wymer and
Lloyd 1996, Hellmann and Wernicke 1998, Schröder et al.
2001). This raises the intriguing question as to what extent gtubulin-associated nucleation occurs not only in cycling but
also in differentiating plant cells. To approach this issue, we
Key words: Cytoskeleton — Gene expression — Hordeum
vulgare — Nicotiana tabacum — Phylogeny — g-tubulin.
Abbreviations: MTOC, microtubule organizing centre; pfu,
plaque forming unit.
Introduction
Successions of diverse arrays of microtubules drive crucial steps in cell division and differentiation in higher plants.
The major microtubule patterns include preprophase bands
related to determining the orientation of the future cell division
plane, spindles for separating chromatids or chromosomes,
phragmoplast microtubules involved in building the new cell
plate and various cortical arrays associated with wall pattern1
Corresponding author: E-mail, [email protected]; Fax, +49-6131-3923075.
224
g-Tubulin in barley and tobacco
225
Fig. 1 Comparison of predicted complete amino acid sequences of the g-tubulin cDNAs derived from barley (Hordeum vulgare: H. v.) and
tobacco (Nicotiana tabacum: N. t.), including the first known g-tubulin sequence (Aspergillus nidulans: A. n.). Sequences were aligned and displayed using the software programmes ClustalX and GeneDoc, respectively. The barley sequence was taken as the reference. Dots indicate
sequence homologies; dashes indicate gaps included for optimal alignment, # indicate end of protein sequence corresponding to translational stop
codon. Arrows mark the locations of the primers initially used for amplifying conserved open reading frames and thus probes.
tested a dicotyledonous and a monocotyledonous species,
tobacco and barley, respectively, for the presence of expressed
g-tubulin genes. cDNA sequences were isolated and compared
to known g-tubulins to determine potential plant-specific features. In addition, Northern blot experiments were performed to
reveal general expression patterns during early development of
leaves, as model systems covering the phases of cell division
and differentiation.
Results
Isolation and characterisation of the C-tubulin cDNA sequences
The potentially g-tubulin-specific PCR primer-sets applied
in barley yielded two fragments, 621 and 656 bp in length,
respectively. Both fragments were 100% identical in their overlapping sequence and database alignment showed highest
homologies to the already known sequence of rice (Oryza
sativa) g-tubulin. Both sequences were used as conserved gtubulin-specific probes in further experiments with barley.
Probing of cDNA libraries yielded several positive clones. In a
library derived from the meristematic base of the leaf, i.e. the
basal 5 mm, 21 clones were found in a screening of 5.5´105
plaque-forming units (pfu). The yield was one order of magnitude lower when a library derived from an only slightly more
advanced developmental stage, 5–10 mm, was tested. The clones
appeared to be identical with one exception. There were two
variants with regard to the length of the 3¢-untranslated region
(UTR), 456 bp and 285 bp, respectively. Southern blot analysis
of genomic DNA digested with restriction enzymes (EcoRI,
HindIII, XbaI, data not shown) indicated the presence of only
one g-tubulin gene. The deduced amino acid sequence of the
complete open reading frame is shown in Fig. 1, for nucleotide
sequence see accession no. AJ272313, GenBank data base.
Fig. 2 Phylogenetic tree for predicted amino acid sequences of barley and tobacco and all the other plant g-tubulins known so far and the
human g-tubulin, with Aspergillus nidulans as outgroup sequence. The
tree was derived using the neighbour-joining method (PHYLIP 3.5c
software package: J. Felsenstein, University of Washington, Seattle,
U.S.A.). Numbers at nodes represent the confidence level of 100 bootstrap samples. The horizontal bar indicates ‘percentage accepted mutations’ (PAM) distance. Non-barley or -tobacco sequences were taken
from the Swissprot database.
In tobacco, application of the more degenerate primers
resulted in the amplification of a 845 bp fragment that exhibited highest homology to the already known g-tubulin
sequences of Arabidopsis. A cDNA library derived from a
226
g-Tubulin in barley and tobacco
Fig. 3 Northern blots of barley, aligned with a diagram of the leaf
and a bar graph showing the mitotic indices (mean derived from four
leaves) of the corresponding leaf regions. No mitotic activity was
observed beyond the region to 9 mm from the leaf base. Twelve mg
RNA were blotted per sample. Please note that the intensities of the
signals represent relative values. No conclusions can be drawn as to
absolute ratios between a- and g-tubulin levels.
Fig. 4 Northern blots of various tobacco tissues (numbered consecutively). The sketch indicates the shoot tip, including the successive
leaves, from which the various RNA samples were derived. The bar
graph shows the mitotic indices of the corresponding tissues (mean
derived from six shoots). Twelve mg RNA were blotted per sample.
highly meristematic cell suspension culture (mitotic index >6)
yielded 17 positive clones after screening of only 4.2´104 pfu
with the potentially tobacco g-tubulin-specific, RT-PCR derived
probe. The predicted amino acid sequence of a full length
clone, aligned with the barley sequence and with the first gtubulin sequence identified (Aspergillus nidulans), is shown in
Fig. 1; for complete nucleotide sequence see accession no.
AJ278739. A phylogenetic sequence analysis including all
known deduced amino acid sequences for g-tubulin is shown in
Fig. 2. These sequences not only include tobacco (Nicotiana
tabacum) and barley (Hordeum vulgare) and the other
angiosperms, i.e. Arabidopsis thaliana, maize (Zea mays) and
rice (Oryza sativa) (Liu et al. 1994, Lopez et al. 1995, Kim et
al. 1999), but also the green alga Chlamydomonas reinhardtii
(Silflow et al. 1999), the moss Physcomitrella patens (GenBank accession no AF142098) and the fern Anemia phyllitidis
(Fuchs et al. 1993).
Northern blot analysis
A scheme representing the barley leaf analysis is depicted
in Fig. 3. Mitotic indices were determined in 1 mm long intervals to get better temporal/spatial resolution, starting with the
leaf base. Meristematic activity was restricted to the basal
9 mm of the leaf, with highest rates in the first 5 mm. No
mitotic figures were found beyond this region. Samples taken
in regular intervals of 5–10 mm (compare Fig. 3) were subjected to Northern blot analysis, starting at the base and extending up to 70 mm, thus covering the meristem and the whole
area of cell expansion and shaping. Samples were probed with
conserved barley g-tubulin sequences. a-Tubulin RNA expression was taken as an appropriate and convenient reference
(Schröder et al. 2001). There appeared to be a distinct correlation between developmental stage and tubulin expression. The
a-tubulin expression pattern was similar to that described
recently (Schröder et al. 2001), with highest levels found in the
g-Tubulin in barley and tobacco
227
Fig. 5 g-Tubulin sequence comparison across different phyla and/or taxa. Only the domain referred to as g-Peptide B by Burns (1995) is shown,
including the conserved flanking regions. The conserved, apparently multicellular plant-specific motif is boxed. Note that only ‘conventional’ gtubulins (Oakley 2000) are compared, i.e. Saccharomyces and Caenorhabditis sequences are not included. Sequences were initially aligned with
the software ClustalX and then manually amended to maximize homologies and displayed using the GeneDoc program. Non-barley or -tobacco
sequences were taken from the Swissprot data base.
cell expansion and shaping region. However, highest g-tubulin
RNA steady-state levels were found in the meristematic basal
5 mm of the leaf, followed by a steady decline in more
advanced regions. Densitometric analysis of the blots revealed
that steady-state RNA g-tubulin levels in early post-mitotic tissue (10–15 mm distance from the leaf base) dropped to ca 30%
of that found in the most basal, meristematic section (0–5 mm).
Nevertheless, there was still a ca 10% level in the clearly nonmeristematic, cell-shaping region of 20–25 mm distance from
the meristematic base. Incidentally, no differences in the
expression patterns were found between the two g-tubulins
showing different lengths in the 3¢-UTR.
Tobacco leaves do not exhibit the stringent developmental gradient found in grass leaves. To overcome this limitation,
whole leaves of various developmental stages had to be taken,
starting with the shoot tip. A diagram of the corresponding
tobacco shoot with its succession of leaves is shown in Fig. 4.
The youngest sample taken consisted of the shoot apical meristem including the highly meristematic ensheathing leaf primordia with a maximum length of ca 2 mm (1). The next older
stage was a rapidly expanding leaf of ca 30 mm length (2), followed by still growing leaves of ca 60 and 100 mm (3 and 4,
respectively). The next leaf (5) in the succession was ca
180 mm long and almost fully expanded. The oldest leaves
included were ca 200 mm in length (6) and just fully expanded.
There was a steady decline in the mitotic indices with increasing leaf age. No apparent mitotic activity was found in the
almost fully expanded leaf 5 and beyond (Fig. 4). The g-tubulin
expression pattern detected was basically similar to that
observed in barley with highest steady-state g-tubulin RNA
levels found in the samples with the highest meristematic rates.
The g-tubulin RNA levels decreased more rapidly than those of
a-tubulin. Consequently, a-tubulin expression was still found
in the fully expanded, no longer meristematic leaves, but only
barely detectable traces of g-tubulin RNA.
Discussion
The present study adds two angiosperm species to the still
brief list of plants with known g-tubulin sequences. In fact,
tobacco represents only the second dicotyledonous and barley
the third monocotyledonous species. A phylogenetic analysis
of predicted protein sequences revealed distinct clusters (Fig.
2). The multicellular green land plants were clearly separated
from the unicellular green alga Chlamydomonas. The archegoniates Anemia and Physcomitrella formed a cluster, as did
the angiosperms. Within the angiosperms a cluster corresponded to dicots and another to monocots or, more precisely,
to grasses. A comprehensive sequence alignment including gtubulin sequences of various organisms confirmed relatively
conserved domains, but also highly variable regions, such as
the C-terminus. Most interestingly, there were particular gtubulin-specific, closely linked domains, referred to as g-Peptide A and B by Burns (1995) that revealed conserved motifs,
apparently specific to multicellular plants. In g-Peptide A it was
the motif VERQ(A/V)N (data not shown, compare Burns
1995); g-Peptide B exhibited the motif SYAR(T/N)KEASQAK
(Fig. 5). Both motifs are flanked by sequences that are highly
conserved across most phyla and/or taxa. It appears that the
corresponding domains are exposed in the folded protein
228
g-Tubulin in barley and tobacco
(Burns 1995), permitting interactions with associated proteins,
e.g. in the ring-forming complexes (Inclán and Nogales 2001).
Perhaps the variant motifs are involved in allowing g-tubulin to
perform its particular functions in multicellular land plants
lacking centrosomes. In this context it should be noted that the
flagellated unicellular green alga Chlamydomonas exhibited a
different peptide. It will be very worthwhile to elucidate to
what extent the differences in g-tubulin sequences observed
reflect simple phylogenetic distances or actual functional specialisation (see also Oakley 2000).
Reports describing attempts to localize g-tubulin in plants
by immunofluorescence microscopy suggest a surprisingly dispersed distribution of g-tubulin, almost mirroring every microtubule pattern (e.g. Joshi and Palevitz 1996, Canaday et al.
2000, Panteris et al. 2000). Most data reported were derived
from meristematic tissues or cell cultures, but apparent colocalization has also been found in cortical arrays of interphase
cells (e.g. Canaday et al. 2000). Interestingly, a more localised
distribution of g-tubulin has been implicated in the organisation
of cortical microtubules of post-mitotic, differentiating guard
cells, associated with wall deposition and cell shaping
(McDonald et al. 1993). These data indicate at least presence, if
not function of g-tubulin throughout all phases of microtubule
occurrence. It remains to be seen whether the apparent decoration of plant microtubules with g-tubulin is actually indicative
of highly dispersed nucleation sites or a result of further functions or features (Joshi and Palevitz 1996, Canaday et al. 2000,
Panteris et al. 2000). An analysis of gene expression at the
RNA level could give some additional clues concerning expression and perhaps evolution of function of g-tubulin in plants.
Our Northern blot experiments indicate that g-tubulin gene
expression in higher plants coincides with mitotic, rather than
with post-mitotic activities of the microtubular cytoskeleton.
This was particularly obvious in the barley leaf with its stringent developmental gradient typical of monocotyledonous
grass species, even though steady-state levels of g-tubulin RNA
did not drop as dramatically as apparent mitotic rates. The barley leaf experimental system permitted discrimination between
timing of mitotic activity and of post-mitotic growth, much better than in the dicotyledonous tobacco. In grasses, such as barley, intricate rearrangements of microtubular arrays occur in
developing leaves after completion of the last mitotic cycle and
result in the formation of complex cell shapes in the terminally
differentiating leaf tissue (Wernicke and Jung 1992). These
highly synchronous post-mitotic rearrangements correlated
with a transient increase in tubulin levels in early post-mitotic
cells (compare Fig. 3) and with differential expression of members of the a-tubulin gene family in a characteristic fashion,
indicating that they are not simply mediated by shifting whole,
stable microtubules (Hellmann and Wernicke 1998, Schröder et
al. 2001). The g-tubulin expression data presented here could
suggest that the post-mitotic changes in microtubule arrays are
less dependent on concurrent g-tubulin gene expression at the
RNA level than mitotic stages. It will be worthwhile to eluci-
date whether the residual, but distinct g-tubulin RNA levels
observed in post-mitotic tissues are of significance in this context. We do not wish to over-interpret our current results. In
order to be able to draw further conclusions data need to be
quantified, but then additional cell-cycle- and differentiationspecific parameters, not a mitotic index alone, should be taken
as a reference. Nevertheless, the data as shown here reveal the
intriguing fact that g-tubulin RNA expression predominates in
mitotic higher plant cells, despite the absence of centrosomes.
Yet, gene activity was still apparent in post-mitotic tissues, if
the presence of RNA is a valid indicator. Perhaps the establishment of the conserved novel motifs described played a crucial
role within the context of loss of centrosomes and gain of
novel, plant specific functions during phylogeny.
Materials and Methods
Plant material
Barley plants (Hordeum vulgare L., cv Igri, seeds kindly provided by Ackermann Co., Irlbach, Germany) were grown in vermiculite in a growth cabinet under standardised conditions, as described
(Hellmann et al. 1995). Second foliage leaves (100–120-mm long) were
selected for analysis. This size was reached ca 9 d after sowing. Leaves
were dissected transversely into sections of defined length (compare
Fig. 3), starting at the base. The distance from the base strictly correlated with the developmental state of the tissue because of the developmental gradient along the leaf, characteristic of growing grass leaves.
Tobacco plants (Nicotiana tabacum L. cv Petit Havana, SR I)
were grown from seeds in a greenhouse. The upper part of shoots comprising the apical meristem and a succession of leaves ranging from
young primordia to fully expanded leaves (compare diagram in Fig. 4)
were excised from the plants about 10 weeks after sowing. The leaf
lamina was processed after removal of the midrib, when possible. A
rapidly proliferating friable cell suspension established from the same
tobacco line several years ago was grown on a rotary shaker in
Murashige and Skoog (1962) liquid medium supplemented with 5 mM
2,4-dichlorophenoxyacetic acid.
Mitotic indices were determined after fixing of tissues in ethanol/
acetic acid and staining of chromatin in aceto-orcein or -carmin. A
minimum of 1,000 cells was counted in each tissue sample. The data
shown in the graphs represent means derived from 4–6 samples. No
standard deviations have been shown, to avoid a degree of precision
not intended. The only purpose of the counts was to show general tendencies and, particularly, to delimit the phase of meristematic activity
from that of post-mitotic growth.
RNA isolation and establishment of cDNA-libraries
RNA was isolated using guanidine hydrochloride/LiCl as
described recently (Schröder et al. 2001) or a commercial product
(peqGold RNAPure, peqlab Biotechnologie GmbH, Erlangen,
Germany). RNA concentrations were determined photometrically.
For construction of cDNA-libraries, poly(A)+-RNA was recovered from total RNA by oligo(dT)-cellulose column chromatography
(Stratagene, Heidelberg, Germany) according to Sambrook et al.
(1989) and primed with an oligo(dT)-tailed linker-primer for cDNA
preparation. cDNA libraries were established using the Uni-ZAP™
XR- or ZAP Express™ vector-systems (Stratagene, Heidelberg,
Germany). Positive clones were identified by plaque lift hybridisation
(Benton and Davies 1977) and sequenced by the dideoxy chain termination method (Sanger et al. 1977).
g-Tubulin in barley and tobacco
Probes for cDNA library screening
Total RNA preparations were subjected to RT-PCR with sets of
degenerate primers. In barley, RNA was isolated from the basal 10 mm
of the young leaf, comprising the highly meristematic base and the
early elongation zone. The primers were designed after taking into
account known maize g-tubulin sequences. One downstream primer
(GD3: GCC AAC CAG ATC ATT GTT C) was combined with two
different upstream primers covering different regions (GU1: TCG
AGG ACT TCG C(C/T)A CTC A; GU2: GA(C/T) GTC TTC TTT
TAC CAG G). In tobacco, RNA was isolated from a highly meristematic cell suspension. The primers were designed with a higher degree
of degeneracy, because the more specific primers used for barley
yielded no signals. Upstream primer, GU2-DEG: GA(C/T) GTI TT(C/T)
TT(C/T) TA(C/T) CA(A/G) GC; downstream primer according to
Fuchs et al. (1993), sequence information kindly confirmed by Dr. B.
Moepps. The positions of the barley and tobacco primers are indicated
in Fig. 1. Annealing temperatures were determined according to
Sambrook et al. (1989). PCR was performed using the Primus 25
thermal cycler (MWG Biotech, Ebersberg, Germany).
The PCR-fragments obtained were electrophoresed and cloned
into pBluescript vectors (Stratagene, Heidelberg, Germany). Probes for
screening cDNA libraries were labeled with [a-32P]dATP by random
priming (Feinberg and Vogelstein 1983) using the RadPrime DNA
Labeling System (GibcoBRL, Eggenstein, Germany).
Northern blotting
Samples of total RNA were mixed with denaturing buffer and
blotted onto nylon membranes (Hybond-N+, Amersham-Pharmacia,
Freiburg, Germany). Probes used were labeled with [a-32P]dATP by
PCR, similar to Mertz and Rashtchian (1994), in order to obtain selective labeling with high specific activity. Hybridisation was performed
under high-stringency conditions at 65°C according to Hellmann et al.
(1995). After hybridisation the membranes were washed twice in
buffer containing 5% SDS and twice in 1% SDS buffer. The blots were
exposed to imaging plates (BAS-MP 2025P, Fujifilm, Japan), that
were scanned in a phospho-imager (BAS 1500, Raytest, Straubenhardt, Germany). The strongest signals were set to 100% on a linear
gray scale for both, a- and g-tubulin. Changes in relative levels of gtubulin RNA were directly referred to those of a-tubulin, because it
would be difficult to find a constitutively expressed control in the rapidly developing tissues analysed. In present work it was not intended
to determine absolute ratios of concentration between a- and g-tubulin;
see Schröder et al. (2001) for a method to determine such ratios.
Acknowledgments
We wish to thank our colleagues for valuable discussion and/or
assistance, especially Silke Heichel and Markus Kriebisch.
References
Benton, W.D. and Davies, R.M. (1977) Screening l-gt recombinant clones by
hybridization to single plaques in situ. Science 196: 180.
Burns, R.G. (1995) Analysis of the g-tubulin sequences: Implications for the
functional properties of g-tubulin. J. Cell Sci. 108: 2123–2130.
Canaday, J., Stoppin-Mellet, V., Mutterer, J., Lambert, A.M. and Schmit, A.C.
(2000) Higher plant cells: Gamma-tubulin and microtubule nucleation in the
absence of centrosomes. Microscopy Res. Technol. 49: 487–495.
Feinberg, A.P. and Vogelstein, B. (1983) A technique for radiolabeling DNA
restriction endonuclease fragments to high specific activity. Anal. Biochem.
132: 6–13.
229
Fuchs, U., Moepps, B., Maucher, H.P. and Schraudolf, H. (1993) Isolation, characterization and sequence of a cDNA encoding gamma-tubulin protein from
the fern Anemia phyllitidis L. Sw. Plant Mol. Biol. 23: 595–603.
Hellmann, A., Meyer, C.U. and Wernicke, W. (1995) Tubulin gene expression
during growth and maturation of leaves with different developmental patterns. Cell. Motil. Cytoskeleton 30: 67–72.
Hellmann, A. and Wernicke, W. (1998) Changes in tubulin protein expression
accompany reorganization of microtubular arrays during cell shaping in barley leaves. Planta 204: 220–225.
Hush, J.M., Wadsworth, P., Callaham, D.A. and Hepler, P.K. (1994) Quantification of microtubule dynamics in living plant cells using fluorescence redistribution after photobleaching. J. Cell Sci. 107: 775–784.
Hyman, A. and Karsenti, E. (1998) The role of nucleation in patterning microtubule networks. J. Cell Sci. 111: 2077–2083.
Inclán, Y.F. and Nogales, E. (2001) Structural models for the self-assembly and
microtubule interactions of g-, d- and e-tubulin. J. Cell Sci. 114: 413–422.
Joshi, H.C. and Palevitz, B.A. (1996) Gamma tubulin and microtubule organization in plants. Trends Cell Biol. 6: 41–44.
Kim, Y.K., Cha, Y.K., Jun, H.Y., Kim, J.D., Choi, J.S., Kim, H.R. and Han, L.S.
(1999) Nucleotide sequence of a cDNA (OstubG2) (accession no. AF036957)
encoding a gamma-tubulin in the rice plant (Oryza sativa). Plant Physiol.
121: 1383.
Liu, B., Joshi, H.C., Wilson, T.J., Silflow, C.D., Palevitz, B.A. and Snustad, D.P.
(1994) g-Tubulin in Arabidopsis – gene sequence, immunoblot, and immunofluorescence studies. Plant Cell 6: 303–314.
Liu, B., Marc, J., Joshi, H.C. and Palevitz, B.A. (1993) A g-tubulin-related
protein associated with the microtubule arrays of higher plants in a cell cycledependent manner. J. Cell Sci. 104: 1217–1228.
Lopez, I., Khan, S., Sevik, M., Cande, W.Z. and Hussey, P.J. (1995) Isolation of
a full-length cDNA encoding Zea mays g-tubulin. Plant Physiol. 107: 309–
310.
McDonald, A.R., Liu, B., Joshi, C. and Palevitz, B.A. (1993) g-Tubulin is associated with a cortical-microtubule-organizing zone in the developing guard
cells of Allium cepa L. Planta 191: 357–361.
Mertz, L.M. and Rashtchian, A. (1994) Nucleotide imbalance and polymerase
chain reaction: effects on DNA amplification and synthesis of high specific
activity radiolabeled DNA probes. Anal. Biochem. 221: 160–165.
Moore, R.C., Zhang, M., Cassimeris, L. and Cyr, R.J. (1997) In vitro assembled
plant microtubules exhibit a high state of dynamic instability. Cell. Motil.
Cytoskeleton 38: 278–286.
Moritz, M., Braunfeld, M.B., Guénebaut, V., Heuser, J. and Agard, D.A. (2000)
Structure of the g-tubulin ring complex: a template for microtubule nucleation. Nature Cell Biol. 2: 365–370.
Murashige, T. and Skoog, F. (1962) A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15: 473–497.
Oakley, B.R. (2000) g-Tubulin. Curr. Top. Dev. Biol. 49: 27–54.
Oakley, C.E. and Oakley, B.R. (1989) Identification of g-tubulin, a new member
of the tubulin superfamily encoded by mipA gene of Aspergillus nidulans.
Nature 338: 662–664.
Panteris, E., Apostolakos, P., Graf, R. and Galatis, B. (2000) Gamma-tubulin
colocalizes with microtubule arrays and tubulin paracrystals in dividing vegetative cells of higher plants. Protoplasma 210: 179–187.
Sambrook, J., Fritsch, S. and Maniatis, T. (1989) Molecular Cloning – A
Laboratory Manual. Cold Spring Harbour Laboratory Press, New York.
Sanger, F., Nicklen, S. and Coulsen, A.R. (1977) DNA sequencing with chain
termination inhibitors. Proc. Natl. Acad. Sci. USA 69: 1408–1412.
Schröder, J., Stenger, H. and Wernicke, W. (2001) a-Tubulin genes are differentially expressed during leaf cell development in barley (Hordeum vulgare L.).
Plant Mol. Biol. 45: 723–300.
Silflow, C.D., Liu, B., LaVoie, M., Richardson, E.A. and Palevitz, B.A. (1999)
g-Tubulin in Chlamydomonas: Characterization of the gene and localization
of the gene product in cells. Cell. Motil. Cytoskeleton 42: 285–297.
Vaughn, K.C. and Harper, J.D.I. (1998) Microtubule organizing centers and
nucleating sites in land plants. Int. Rev. Cytol. 181: 75–149.
Wernicke, W. and Jung, G. (1992) Role of cytoskeleton in cell shaping of developing mesophyll of wheat (Triticum aestivum L.). Eur. J. Cell Biol. 57: 88–94.
Wymer, C. and Lloyd, C. (1996) Dynamic microtubules: implications for cell
wall patterns. Trends Plant Sci. 1: 222–228.
(Received August 21, 2001; Accepted December 12, 2001)