Báo cáo khoa học: Different roles of two c-tubulin isotypes in the cytoskeleton of the Antarctic ciliate Euplotes focardii Remodelling of interaction surfaces may enhance microtubule nucleation at low temperature doc

Different roles of two c-tubulin isotypes in the
cytoskeleton of the Antarctic ciliate Euplotes focardii
Remodelling of interaction surfaces may enhance microtubule
nucleation at low temperature
Francesca Marziale
1
, Sandra Pucciarelli
1
, Patrizia Ballarini
1
, Ronald Melki
2
, Alper Uzun
3
,
Valentin A. Ilyin
3
, H. W. Detrich III
3
and Cristina Miceli
1
1 Dipartimento di Biologia Molecolare, Cellulare e Animale, University of Camerino, Italy
2 Laboratoire d’Enzymologie et Biochimie Structurales, Centre National de la Recherche Scientifique, Gif-sur-Yvette, France
3 Department of Biology, Northeastern University, Boston, MA, USA
Microtubule assembly in metazoan cells is nucleated
by organizing centers, which include centrioles, basal
bodies, and other structures. Mitotic centrosomes con-
tain a pair of centrioles and associated pericentriolar
material, whereas basal bodies recruit other accessory
structures [1,2]. Both centrioles and basal bodies
require c-tubulin, the ubiquitous third member of the

‘tubulin superfamily’ [3–5], for their assembly and
maintenance [6–8], and for their capacity to nucleate
microtubules [9]. This tubulin variant associates with
Keywords
microtubule nucleation; molecular cold-
adaptation; psychrophilic microorganism;
quantitative PCR; tubulin genes
Correspondence
C. Miceli, Dipartimento di Biologia
Molecolare, Cellulare e Animale, University
of Camerino, Via Gentile III da Varano,
62032 Camerino (MC), Italy
Fax: +39 0737 40 32 90
Tel: +39 0737 40 32 55
E-mail:
(Received 22 July 2008, revised 27 August
2008, accepted 4 September 2008)
doi:10.1111/j.1742-4658.2008.06666.x
c-Tubulin belongs to the tubulin superfamily and plays an essential role in
the nucleation of cellular microtubules. In the present study, we report the
characterization of c-tubulin from the psychrophilic Antarctic ciliate Eupl-
otes focardii. In this organism, c-tubulin is encoded by two genes, c-T1 and
c-T2, that produce distinct isotypes. Comparison of the c-T1 and c-T2 pri-
mary sequences to a Euplotes c-tubulin consensus, derived from mesophilic
(i.e. temperate) congeneric species, revealed the presence of numerous
unique amino acid substitutions, particularly in c-T2. Structural models of
c-T1 and c-T2, obtained using the 3D structure of human c-tubulin as a
template, suggest that these substitutions are responsible for conformational
and ⁄ or polarity differences located: (a) in the regions involved in longitudi-
nal ‘plus end’ contacts; (b) in the T3 loop that participates in binding GTP;

and (c) in the M loop that forms lateral interactions. Relative to c-T1, the
c-T2 gene is amplified by approximately 18-fold in the macronuclear gen-
ome and is very strongly transcribed. Using confocal immunofluorescence
microscopy, we found that the c-tubulins of E. focardii associate throughout
the cell cycle with basal bodies of the non-motile dorsal cilia and of all of
the cirri of the ventral surface (i.e. adoral membranelles, paraoral mem-
brane, and frontoventral transverse, caudal and marginal cirri). By contrast,
only c-T2 interacts with the centrosomes of the spindle during micronuclear
mitosis. We also established that the c-T1 isotype associates only with basal
bodies. Our results suggest that c-T1 and c-T2 perform different functions
in the organization of the microtubule cytoskeleton of this protist and are
consistent with the hypothesis that c-T1 and c-T2 have evolved sequence-
based structural alterations that facilitate template nucleation of microtu-
bules by the c-tubulin ring complex at cold temperatures.
Abbreviations
qPCR, quantitative PCR; RATE, rapid amplification of telomeric ends; TuRC, tubulin ring complex.
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5367
other proteins to form two macromolecular structures,
the c-tubulin small complex, which possesses a weak
microtubule nucleating activity [10,11], and the c-tubu-
lin ring complex (TuRC) [12], which nucleates
strongly. c-TuRC resembles a lock washer and is con-
sidered to be the fundamental unit required for micro-
tubule nucleation. Two models have been proposed to
explain microtubule nucleation by c-TuRC: (a) the
‘protofilament’ model, in which the c-tubulin subunits
of c-TuRC associate longitudinally with ab-tubulin
dimers [13], and (b) the ‘template’ model, in which the
c-TuRC ring mimics the end of a microtubule, and
c-tubulin interacts both longitudinally and laterally

with a-tubulin but only laterally with b-tubulin [14,15].
Microtubule assembly is entropically driven, pre-
dominantly via hydrophobic interactions, and therefore
is sensitive to environmental temperature both in vitro
and in vivo [16,17]. The ab-tubulin dimers of mam-
mals, for example, form microtubules in vitro at tem-
peratures near 37 °C, and these polymers dissociate at
low temperature (4 °C) to yield tubulin dimers and
ring-shaped oligomers [18–20]. Ectothermic (cold-
blooded) Antarctic fishes, by contrast, possess tubulins
that polymerize at temperatures as low as )1.8 °C,
which is the freezing point of their chronically cold
marine habitat [16,17]. Detrich and colleagues have
shown that thermal compensation of microtubule
assembly and dynamics in these fishes results from the
evolution of structural changes intrinsic to the a- and
b-tubulins [21–24].
The nucleation of cytoplasmic microtubules by cen-
trosomes requires productive binding reactions
between c-tubulin and the ab-tubulin dimer, but the
molecular alterations that conserve nucleation in cold-
living organisms have not been studied. Data indirectly
relevant to temperature compensation of microtubule
nucleation were obtained from alanine-scanning muta-
genesis of the c-tubulins of Tetrahymena thermophila
[25] and Aspergillus nidulans [26]. Substitution of ala-
nine at sites in the lateral surfaces (the H3 helix and
the M loop) of these c-tubulins causes cold-sensitivity
of cell growth and ⁄ or loss of basal bodies [25,26]. In
light of this evidence, we propose that the capacity of

c-tubulin to perform efficient microtubule nucleation
at cold temperatures reflects evolved molecular altera-
tions to its interaction surfaces.
Psychrophilic ciliated protozoa are uniquely suited
to an investigation of this issue. As single cells, ciliates
are directly exposed to environmental factors through-
out their life cycle, and modifications of the primary
sequences of many of their proteins are likely to reflect
adaptive mutations that increase the fitness of the
organism at cold temperatures. In ciliates, microtubule
nucleation is promoted mainly by basal bodies, which
are positioned precisely in organized rows in the
somatic cell cortex and in the oral apparatus [8]. The
assembly and maintenance of basal bodies were both
shown to require c-tubulin [7,8].
The ciliate Euplotes focardii, which is endemic to Ant-
arctic coastal seawaters, shows strictly psychrophilic
phenotypes, including optimal survival and multiplica-
tion rates at 4–5 °C [27], the lack of a transcriptional
response of the Hsp70 genes to thermal shock [28], and
modifications in the primary structures of the a- and
b-tubulin [29–31] and of the proteins that form the ribo-
somal stalk [32]. In the present study, we characterized
the two c-tubulin isotypes, c-T1 and c-T2, of E. focardii ,
model their 3D structures, and examined their differen-
tial expression and cellular localization. We suggest that
novel amino acid substitutions located at the plus ends,
near the GTP-binding sites, and within the M loops of
the E. focardii c-tubulins, preserve their microtubule-
nucleating activities at cold temperatures and ⁄ or confer

different functions on the two isotypes.
Results
Sequence analysis of E. focardii c-tubulin genes
Two c-tubulin genes (nanochromosomes), designated
c-T1 (1623 bp; GenBank accession number EF189704)
and c-T2 (1619 bp; GenBank accession number
EF189705), were obtained by our rapid amplification
of telomeric ends (RATE)-PCR-based cloning strategy.
The existence of more than two c-tubulin genes in
E. focardii was excluded by restriction analysis of
macronuclear DNA. Figure 1A shows that undiges-
ted macronuclear DNA gave a single band of approxi-
mately 1.6 kb (lane 1) when hybridized at low
stringency to a probe derived from the c-T2 gene.
Co-digestion of macronuclear DNA by EcoRI and
HindIII (lane 2) gave strongly hybridizing fragments of
approximately 640, 480, and 300 bp, and weakly
hybridizing bands of approximately 750 and 200, con-
sistent with the lengths and restriction maps of the two
c-tubulin nanochromosomes (Fig. 1B). The restriction
maps and relative abundances of the DNA fragments
suggest that the c-T2 nanochromosome is amplified to
a greater extent than the c-T1 nanochromosome. Both
isotypes are expressed, as shown by the recovery of
distinct c-T1 and c-T2 cDNAs of approximately
1.4 kb. Furthermore, northern blot analysis of mRNA
extracted from exponentially growing E. focardii cells,
when hybridized at low stringency to the c-T2 probe,
indicated that the two mRNAs were comparable in
size (1.4 kb; not shown).

c-Tubulin isotypes in E. focardii F. Marziale et al.
5368 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
The coding sequences of the E. focardii c-T1 and
c-T2 nanochromosomes were interrupted by two
introns located in identical positions (Fig. 1B). The first
intron included nucleotides 50–96 and the second intron
included nucleotides 210–253 in each gene. Excluding
introns and stop codons, the c-T1 and c-T2 coding
regions were each 1383 bp in length and predicted
proteins of 461 amino acids. The nucleotide sequence
identity between c-T1 and c-T2 was 94.6%. Two
in-frame UGA codons, which are known to code for
cysteine in other Euplotes species [33,34], were present at
residue positions 109 and 185 in each of the genes.
Comparative structural modelling of E. focardii
c-tubulins to human c-tubulin
The 3D structures of the E. focardii c-tubulins were
modeled comparatively with respect to human c-tubu-
lin [35]. The predicted structures of c-T1 and c-T2
were remarkably similar to that of the human protein
(Fig. S1).
Structural features of E. focardii c-tubulin
isotypes
Plus ends
The deduced amino acid sequences of the c-T1 and
c-T2 isotypes were aligned with respect to a Euplotes
c-tubulin consensus sequence and mapped onto the
consensus secondary structure of the tubulin mono-
mer [35,36] (Fig. 2). c-T1 and c-T2 were 95.4% iden-
tical in amino acid sequence. The main differences of

the two isotypes compared to the Euplotes c-tubulin
consensus were found in two regions, 390–403 and
70–95 (Fig. 2), both of which are located at the plus
end (Fig. 3). In the former, c-T2 contained several
polar-for-charged substitutions (K394S, R395N,
D396N, and K403Q) with respect to the consensus
(consensus residue ⁄ residue position ⁄ c-T2 residue).
c-T1 displayed substitutions of bulky residues with
respect to the consensus sequence (T391I, K394R;
consensus ⁄ position ⁄ c-T1), reciprocal changes of polar
and charged amino acids (D396N, N400D), and one
polar-for-hydrophobic alteration (I401N). Notable
amino acid substitutions in the second region (70–95)
of c-T1 and c-T2 with respect to the Euplotes consen-
sus included the V of c-T1 and K of c-T2 for G at
position 76, A of c-T1 ⁄ c-T2 for the consensus P at
position 81, G for S at position 84, F for Y at posi-
tion 92, and S for A at position 94. Together, these
results show that the plus-end surfaces of the two
E. focardii c
-tubulins have diverged considerably
from those of mesophilic Euplotes species, with an
overall tendency toward greater hydrophobicity. By
contrast, very few changes were observed in sequences
that contribute to the c-tubulin minus end (Figs 2
and 3).
Isotypic substitutions
E. focardii c-T1 and c-T2 differed considerably
between themselves at their plus ends. Major residue
changes included R72G, V76K, R394S, R395N,

Y398F, D400T, N401T, and K403Q (c-T1 ⁄ residue
position ⁄ c-T2). This suite of residue substitutions may
confer unique functions upon each isotype.
Fig. 1. The macronucleus of Euplotes focardii contains two differ-
ent c-tubulin nanochromosomes. (A) Southern blot analysis of the
two c-tubulin genes of E. focardii using the c-T2 gene as probe.
Lane 1, undigested DNA; lane 2, EcoRI- and HindIII-digested DNA.
The sizes (bp) of DNA standards are indicated on the left. The sizes
of the two c-tubulin nanochromosomes (1600 bp) and their diges-
tion products are indicated on the right. (B) Structural features and
EcoRI ⁄ HindIII restriction maps of the E. focardii c-T1 and c-T2 nano-
chromosomes. Coding, noncoding regions, introns, and telomeres
(C
4
A
4
⁄ G
4
T
4
) are indicated in the key.
F. Marziale et al. c-Tubulin isotypes in E. focardii
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5369
Nucleotide-binding sites
Human c-tubulin binds GTP in a plus-end cleft
enclosed by residues G11, Q12,
C13, Q16, G101,
N102, S140, A142, G143, G144, T145, V171, P173,
N207, F225, I228, and N229 (where the residues
shown underlined form main- and ⁄ or side-chain

hydrogen bonds with atoms of the nucleotide) [35].
Fig. 2. Sequence comparisons of Euplotes focardii c-T1 and c-T2 with the Euplotes c-tubulin consensus. The unique substitutions of E. focar-
dii c-T1 and c-T2 are shown as a single-letter code underneath the Euplotes c-tubulin consensus sequence; conserved residues are indicated
by dots. Predicted secondary structural elements, H for helices and S for strands [37], are represented by white cylinders and black arrows,
respectively. T1 to T7 indicate loops that are involved in contacts with the bound GTP [37]. Residues involved in longitudinal contacts at the
‘plus’ and ‘minus’ ends are indicated by ‘+’ and ‘)’, respectively, whereas those involved in the lateral contacts of the H3 and M-loop are
shown by ‘H’ and ‘M’. Regions thought to participate in binding to ab tubulin heterodimers [68] are underlined.
c-Tubulin isotypes in E. focardii F. Marziale et al.
5370 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
These residues are all conserved in the E. focardii
c-tubulins. Near the entrance to the nucleotide pocket
within the H2 helix, c-T2 possessed a striking sub-
stitution at position 72: glycine in place of the
c-T1 ⁄ consensus arginine (Figs 2–4). The presence of
glycine at this position in c-T2 has only been
observed in E. focardii and in the psychrotolerant
Euplotes crassus [34, present study]. By contrast, sub-
stitution of alanine for arginine 72 in the c-tubulins
of T. thermophila and A. nidulans produces a lethal
phenotype [25,26], which suggests that this basic resi-
due is important for c-tubulin function at moderate
temperature. The sequences of the nucleotide-binding
T3 and T5 loops of c-T1 and c-T2 were highly con-
served across all Euplotes species; the former
perfectly, whereas the latter contained a single polar-
for-hydrophobic change, I174N (consensus ⁄ position ⁄
c-T1 and c-T2).
Fig. 3. 3D mapping of the sequence substitutions of Euplotes focardii c-T1 and c-T2 with respect to the Euplotes c-tubulin consensus.
(A, B) c-T1 viewed from the side and from the plus end, respectively. (C, D) c-T2 viewed from the side and from the plus end, respectively.
Ribbon diagrams of c-T1 and c-T2 were obtained by comparative modelling to human c-tubulin using

MODELLER, version 9.1 (http://www.
salilab.org/modeller/) [64]. Residues that distinguish the E. focardii c-tubulins from the Euplotes consensus are shown in red and annotated
as consensus residue ⁄ position ⁄ c-T1 or c-T2 residue. The GTP molecule is shown in green. The plus and minus ends, the H3 helix, and the
M-loop are indicated.
F. Marziale et al. c-Tubulin isotypes in E. focardii
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5371
M loops
The ‘extended’ M loop, which we define as encom-
passing the S7-H9 (M) loop, H9, and the H9-S8
loop, and the H3 surfaces of c-tubulin are involved
in lateral contacts [35–37]. Amino acid substitutions
with respect to the Euplotes consensus were found in
the extended M loop in E. focardii c-T1 and c-T2
(Figs 2 and 3). Two hydrophobic-for-hydrophobic
changes occurred near position 280 (F279L, V282I,
consensus ⁄ position ⁄ c-T1 and c-T2) and c-T1 pos-
sessed an alanine at 280 in place of consensus
threonine. The changes in the H9-H9¢ loop were
more dramatic. Both c-T2 and c-T1 contained pro-
line-for-hydroxyl substitutions (T297P and T303P,
respectively).
Tertiary structural differences between E. focardii
c-T1 and c-T2
Figure 4 shows the superimposition of the 3D struc-
tures of c-T1 and c-T2 from the side and the plus end,
respectively. The comparison demonstrates that the
differences between c-T1 and c-T2 (c-T1 ⁄ residue posi-
tion ⁄ c-T2) mapped largely to exposed areas (plus-end
loops and helices, extended M loop) of the polypep-
tides. The valine at S3 position 93 of c-T2 appears to

confer a conformational change in the adjacent T3 loop
(Fig. 4B, double arrow), which is directly involved in
the formation of the GTP-binding site. The alanine at
280 in c-T1 apparently causes a conformational change
in the M loop (S7-H9), which may influence lateral
interactions (Fig. 4A, double arrow). The substitutions
Fig. 4. Comparison of the tertiary structures of Euplotes focardii c-T1 and c-T2. Ribbon diagrams of the two proteins, obtained by compara-
tive modelling to human c-tubulin using
MODELLER, version 9.1 ( [64], are superimposed to highlight structural
differences. The c-T1 and c-T2 loops are shown in yellow and cyan, respectively. Residue substitutions that differentiate the two E. focardii
c-tubulins are colored violet and designated as c-T1 ⁄ residue position ⁄ c-T2. Notable loop displacements are indicated by double arrows. The
GTP molecule is shown in green. The Mg
2+
ion (blue sphere) is shown bound to the b- and c-phosphates of GTP (dark green). (A) Side view.
(A¢,A¢¢) Show close-up side views (generated using Chimera; [69] of the H9-H9¢ loop (dashed box in A),
which contains a proline at position 303 in c-T1 (A¢) in contrast to the serine of c-T2 (A¢¢). The proline substitution of c-T1 eliminates
the hydrogen bond between Ser303 and Asn205 in c-T2. (B) Plus-end view. (B¢) An enlargement of the nucleotide-binding pocket, shown
boxed in (B). Near the entrance to the pocket, c-T2 contains glycine (cyan) in place of the arginine (yellow) normally found at position 72 in
helix H2.
c-Tubulin isotypes in E. focardii F. Marziale et al.
5372 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
of prolines for threonine at position 297 of c-T2 and for
serine at 303 in c-T1 do not alter significantly the con-
formation of the H9-S8 loop (Fig. 4), although they are
likely to restrict its mobility. However, the Pro303 sub-
stitution of c-T1 eliminates the bent hydrogen bond that
forms between Ser303 and Asn205 in c-T2 (compare
Fig. 4A¢,A¢¢). The cluster of substitutions in H11 and
the H11-H12 loop of the two c-tubulins cause polarity
changes at the plus end (Fig. 4) that would differentiate

the longitudinal interactions formed by c-T1 and c-T2.
Finally, c-T2 possesses a glycine at position 72 in place
of consensus ⁄ c-T1 arginine (Fig. 4B¢). This substitution
may ‘open’ the nucleotide-binding site to facilitate
exchange.
We have not attempted to quantify the loop dis-
placements because the T3 and M loops of the 3.0 A
˚
crystal structure of GTP-bound tubulin are disordered
[35]. Hence, we consider the modeled loop displace-
ments of the E. focardii c-tubulins to be provisional
and to require future validation.
Transcription of the E. focardii c-T1 and c-T2
nanochromosomes
To gain insight into the roles of the E. focardii c-tubu-
lin isotypes, we measured the steady-state levels of
macronuclear mRNAs transcribed from the c-T1 and
c-T2 nanochromosomes of starvation-synchronized
cultures by quantitative PCR (qPCR). During starva-
tion, the transcript levels for both isotypes were low
(Fig. 5A). After feeding, the amounts of c-T1 and
c-T2 mRNAs increased, with the latter being two- to
three-fold higher than the former at 18 h. At 36 h
post-feeding, c-T2 mRNA increased 16-fold relative to
its abundance at 18 h (53-fold increase with respect
to t = 0 h), whereas the level of the c-T1 transcript
remained unchanged. Ninety-eight percent of the cells
were undergoing mitosis ⁄ cytokinesis at this time (as
determined by counting of cells using a stereomicro-
scope). By 54 h, c-T2 mRNA returned to a value simi-

lar to that at 18 h, whereas the amount of the c-T1
transcript was comparable to that observed at 18 and
36 h. Thus, the amount of the c-T2 transcript varies
widely during the cell cycle, whereas the c-T1 mRNA
is expressed at low, almost constant levels.
The disparity between c-T1 and c
-T2 transcript lev-
els could result from differential amplification of the
corresponding macronuclear nanochromosomes, as has
been reported for other Euplotes genes [38], from dif-
ferent rates of transcription initiation and elongation
between the two genes, and ⁄ or from variation in the
rates of degradation of the two messages. To test the
first hypothesis, the gene copy number of c-T1 and
c-T2 was estimated by qPCR. c-T1 and c-T2 nano-
chromosomes were present at approximately 175 and
3600 copies per cell, respectively (Fig. 5B). Thus, the
c-T2 template was approximately 21-fold more abun-
dant than the c-T1 template. To evaluate the second
hypothesis, the 5¢ and 3¢ noncoding sequences of the
two c-tubulin genes were compared. Figure 5C shows
that the 5¢-UTR of c-T1 contained the sequence TGA-
TAC ()26 to )21; gray shading), which matches the
consensus sequence for GATA-binding transcription
factors (WGATAR), whereas the c-T2 5¢-UTR pos-
sessed two tandem repeats ()32 to )27, )24 to )19;
gray shading) of the same motif in essentially the same
A B
C
Fig. 5. Macronuclear amplification and tran-

scription of Euplotes focardii c-T1 and c-T2
genes. (A) Cell-cycle-dependence of steady-
state c-T1 and c-T2 mRNA levels deter-
mined by qPCR. Values are the mean ± SD
(n = 4). (B) Determination of macronuclear
gene copy-number of c-T1 and c-T2 by
qPCR. Values are the mean ± SD (n = 4).
(C) Sequences of the 5¢- and 3¢ noncoding
regions of c-T1 and c-T2 putative GATA tran-
scription factor-binding motifs are shown in
gray.
F. Marziale et al. c-Tubulin isotypes in E. focardii
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5373
location. GATA-binding transcription factors are
known to regulate the transcription of some genes in
protists [39]. The 3¢-UTR of c-T2 was three nucleo-
tides shorter than that of c-T1 but, otherwise, these
two sequences were quite similar. We have not yet
investigated the role of message degradation with
respect to the control of c-tubulin transcript abun-
dance. With the latter caveat, we propose that the
quantities of c-T1 and c-T2 mRNAs are regulated, at
least in part, by differential gene amplification and by
the number of GATA-factor promoter motifs.
Distribution of c-tubulins in E. focardii cells
To examine the cellular distribution of c-tubulin, we
used polyclonal anti-(human c-tubulin) serum [40] and
a polyclonal antibody that we prepared against the
most divergent peptide [(390)RIFRRRNAYIDNYK
(403)] of E. focardii c-T1. Figure 6 presents confocal

microscopic images of three E. focardii cells after stain-
ing with antibodies directed against a- and c-tubulins
(cell 1, Fig. 6A–H; cell 2, Fig. 6I–L; cell 3, Fig. 6M–P).
The anti-(human c-tubulin) serum stained all classes of
basal bodies (Fig. 6B,F, red), including those of: (a) the
adoral membranelles that nucleate the microtubules of
the cytostomal ciliature (labeled green by DM1A in
Fig. 6A,C); (b) the paraoral membrane that surrounds
the cytostomal area; (c) the four groups of locomotory
cirri [frontoventral (numbered 1–10), transverse, cau-
dal, and marginal]; and (d) the nonmotile cilia of
the dorsal surface [41], which are arranged in longitudi-
nal rows (kineties). Interestingly, dorsal ciliary micro-
tubules were absent in the equatorial area (Fig. 6E,G),
which suggests that this cell is entering mitosis and that
duplication of basal bodies at the dorsal surface
requires the disassembly of dorsal cilia.
In mitotic E. focardii cells (Fig. 6I–P), the anti-
(human c-tubulin) serum stained newly-formed basal
bodies (Fig. 6J,L, red, arrows) and the poles of the
micronuclear mitotic spindle (Fig. 6J, solid arrow-
head), but macronuclear staining was never observed.
The basal bodies indicated by the upper arrow in
Fig. 6J will form the transverse, caudal and marginal
cirri of the anterior daughter cell, which also inherits
the frontoventral cirri of the parental cell. Conversely,
the basal bodies marked by the lower arrow will pro-
duce the frontoventral cirri of the posterior daughter
cell [42,43] and its transverse, caudal, and marginal
cirri derive from the parent. As division proceeds, the

duplicated basal bodies nucleate new ciliary micro-
tubules of the nascent cirri, as shown by the DMIA
staining in Fig. 6I.
To determine the subcellular localization of c-T1
and c-T2, we attempted to prepare rabbit polyclonal
antibodies specific for the two peptides that clearly dis-
tinguish c-T1 [(390)RIFRRRNAYIDNYK(403)] and
c-T2 [(390)KKLRSNNAFITTYQ(403)]. We obtained
an antibody specific for c-T1. The c-T2 peptide was,
however, not immunogenic. Fig. 6M–O shows that the
anti-c-T1 serum gave staining identical to that
observed with anti-(human c-tubulin), with the excep-
tion that the micronuclear spindle poles were not rec-
ognized. Therefore, we conclude that c-T2, but not c-
T1, participates in the assembly of the mitotic spindle
of E. focardii and that both isotypes are involved in
the nucleation of other microtubule structures.
Finally, we examined the distribution of E. focardii
c-tubulins in total cell extracts and in subfractions
enriched in basal bodies or in micronuclei using anti-
(human c-tubulin) and anti-
c-T1 sera. Figure 7 shows
that the human antibody recognized c-tubulins in all
three samples, whereas the c-T1 antibody gave positive
signals only for the total cell extracts and basal bodies.
These results confirm that c-T2 alone nucleates micro-
tubules in the micronucleus.
Discussion
In the present study, we have shown that the psychro-
philic ciliate E. focardii possesses two c-tubulin genes

Fig. 6. Spatial distribution of c-tubulins in Euplotes focardii cells. Confocal immunofluorescence microscopic images of three E. focardii cells
were recorded after staining with antibodies directed against a- and c-tubulins. Six optical sections, separated by intervals of 1 lm, were col-
lected and merged for each cell ⁄ antigen combination. (A–D) Ventral view of cell 1 in late vegetative stage; (E–H) dorsal view of cell 1; and
(I–L) ventral view of cell 2 in mitosis. The arrowhead in (J) indicates the micronuclear mitotic spindle, and the arrows show the newly-formed
basal bodies. (M–P) Ventral view of cell 3 in mitosis. The arrows in (M) and (O) indicate the micronuclear mitotic spindle. (A–C, E–G, I–K)
Cells were co-stained with mouse monoclonal anti-a-tubulin serum DM1A (Amersham) and rabbit polyclonal anti-(human c-tubulin) serum.
The primary antibodies were detected using Alexa Flour 488 goat anti-(mouse IgG) (green signal indicates microtubules) and Alexa Fluor 594
goat anti-(rabbit IgG) (red signal indicates c-tubulin in basal bodies). (M–O) Cell 3 was stained for microtubules with the primary antibody
DM1A and for c-tubulins with rabbit polyclonal anti-(E. focardii c-T1); secondary antibodies were as before. Co-localization of a- and c-tubulins
is shown by the yellow signals in merged images (C, G, K, O). (D, H, L, P) Black-and-white versions of the merged images are labeled to
identify cytoskeletal structures. am, adoral membranelles; pm, paraoral membrane; 1–10, frontoventral cirri involved in locomotion; tc, trans-
verse cirri; cc, caudal cirri; mc, marginal cirri; mb, microtubule bundles that elongate from the basal bodies of each transverse cirrus into the
cytoplasm. Scale bar = 10 lm.
c-Tubulin isotypes in E. focardii F. Marziale et al.
5374 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
that encode distinct isotypes, c-T1 and c-T2. The
amino acid sequences of the two isotypes have
diverged from those of mesophilic Euplotes species
primarily in two regions that are involved in protein–
protein quaternary interactions: (a) the plus end, which
forms longitudinal contacts, and (b) the extended
ABC D
EF G H
IJK L
MN O P
Mitotic
spindle
Mitotic
spindle
Kinety

Dorsal
Equatorial
area
Newly
formed
basal bodies
cirri
F. Marziale et al. c-Tubulin isotypes in E. focardii
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5375
M loop, which participates in lateral bonding. The
extensive alterations of sequence elements that form
these surfaces are likely to be adaptations that preserve
c-tubulin function at cold temperatures. Moreover,
c-T1 and c-T2 differed substantially in their sequences
at these locations, consistent with the possibility that
the functions of the single c-tubulin found in most
organisms may be partitioned between the two protis-
tan isotypes. Together, our results suggest strongly
that E. focardii has evolved c-tubulins that are able to
nucleate microtubule structures at low temperature
while individually performing specialized subfunctions.
The E. focardii c-tubulin gene family – regulation
of expression
The two c-tubulin genes of E. focardii appear to be a
feature characteristic of this protistan genus. Two
c-tubulin genes have also been reported for
Euplotes octocarinatus [44] and for E. crassus [34]. In
the former case, the c-tubulin genes produce identical
proteins, whereas, in the latter, they encode two differ-
ent isotypes whose functional differences, if any, are

unknown [34].
Transcription of the E. focardii c-T2 gene was
robust and cell-cycle dependent, whereas synthesis of
the c-T1 mRNA occurred at low, almost constant lev-
els. The differential transcription of the two c-tubulin
genes appears to be due to the greater copy number of
the c-T2 nanochromosome in the macronucleus (i.e.
20-fold larger than that of c-T1) and to the duplication
of a GATA-transcription factor binding site in the
c-T2 promoter, although other processes might also be
involved. In multicellular organisms, GATA-binding
factors play critical roles in development, including
cell-fate specification, regulation of differentiation, and
control of cell proliferation and movement [45].
Recently, we have shown that activation of the heat-
shock response in the ciliate T. thermophila requires
GATA motifs, heat-shock transcription elements, and
their cognate transcription factors [39]. Taken together,
our results with protistan genera support the hypothe-
sis that the GATA gene-regulatory system arose early
in metazoan evolution.
Sequence changes in relation to the tertiary and
quaternary structures of the E. focardii c-tubulins –
implications for cold adaptation of tubulins
Although the c-tubulins of E. focardii are strikingly
similar in overall 3D organization to human c-tubu-
lin, the former contain divergent sequence elements
that are likely to affect the mobility of their domains
and their interactions with partner proteins. That the
plus-end surfaces of c-T1 and c-T2 differ physico-

chemically from each other and from that of the
mesophilic Euplotes consensus is clear. The extended
M loops of c-T1 and c-T2 are more hydrophobic
than the Euplotes consensus and contain proline sub-
stitutions whose role may be to constrain the lateral
contact residues to a conformation that is favorable
for formation of the c-TuRC nucleation complex
from multiple c-tubulin small complexes [12].
Similarly, the lateral surfaces of
a-tubulins from
E. focardii and from two psychrophilic algae of the
genus Chloromonas contain hydrophobic substitutions
with respect to the corresponding a-isotypes of tem-
perate congeners [31,46]. Detrich et al. [27] have
shown that a small number of hydrophobic substitu-
tions in Antarctic fish tubulins appear to be impor-
tant for compensatory adaptation of microtubule
assembly at cold body temperatures; one such
change, F200Y (Antarctic fish residue⁄ position ⁄ meso-
philic residue), which is located at the interface
between the nucleotide-binding and intermediate
domains of b-tubulin, clearly affects microtubule
dynamics when mutated in Schizosaccharomyces
pombe [47]. Thus, increased hydrophobicity of tubu-
lins, both at surface interaction sites and at internal
domain interfaces, emerges as a common theme for
psychrophilic organisms.
Sequence alterations near the nucleotide-binding
pocket of E. focardii c-tubulins are also candidates
for adaptive compensation. Amino acid changes

adjacent to the T3 loop and within the T5 loop may
influence the binding affinity and hydrolysis of GTP
and ⁄ or the egress of GDP and P
i
, which may in
turn control the assembly and stability of new basal
bodies [25]. The most striking of these is the G72R
substitution of c-T2 near the entrance to the
AB
Fig. 7. Distribution of c-tubulins in Euplotes focardii nuclei, basal
bodies, and total cell extracts. (A) Total cell extracts (TCE) and sub-
fractions enriched in basal bodies (BB) and in micronuclei (N) were
prepared as described in the Experimental procedures. Western
blots of the extracts and fractions were incubated with (A) poly-
clonal antibodies against human c-tubulin and (B) polyclonal antibod-
ies specific for E. focardii c-T1.
c-Tubulin isotypes in E. focardii F. Marziale et al.
5376 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
nucleotide pocket. This replacement has only been
observed in E. focardii and in the psychrotolerant
E. crassus [34, present study]. By contrast, the similar
substitution of alanine for arginine 72 in the c-tubu-
lins of T. thermophila and A. nidulans yields a lethal
phenotype [25,26]. Together, these results implicate
position 72 as being important for thermal adapta-
tion of c-tubulins and motivate a detailed examina-
tion of the tertiary interactions that surround the
nucleotide-binding site. Finally, GTP hydrolysis may
be also affected by the unique substitution M250L,
which is located adjacent to the motif GxxNxD that

has been proposed to act as a ‘synergistic’ loop in
nucleotide cleavage [37,48,49].
Functions of the E. focardii c-tubulin isotypes
The results obtained in the present study demonstrate
that one or both of the c-tubulins of E. focardii associ-
ate permanently with basal bodies, consistent with
prior observations that c-tubulin is present in these
microtubule organizing centers in the ciliates E. octoca-
rinatus [50], T. thermophila [8], Tetrahymena pyriformis
[51], and Paramecium tetraurelia [52]. Furthermore, we
show that duplication of the basal bodies of the dorsal
surface during cell division in E. focardii is associated,
perhaps causally, with disassembly of the equatorial
cilia and their microtubules. It is tempting to speculate
that the ciliary tubulins are recycled to form the cyto-
plasmic microtubule bundles [42,43] that guide the
positioning of basal bodies in the nascent daughter
cells.
During cell division, the poles of the micronuclear
mitotic spindle of E. focardii stained with an antibody
prepared against human c-tubulin but not with an
antibody specific for the c-T1 isotype, which recog-
nized only basal bodies. Thus, we conjecture that
E. focardii c-T2, whose mRNA levels peak in mitosis,
is the only isotype required for centrosome function in
the closed orthomitosis of the micronucleus. We did
not detect c-tubulin or microtubules in the macronu-
cleus of E. focardii, in contrast to reports that c-tubu-
lin and microtubules are present in the amitotically
dividing macronucleus of T. thermophila [8,53]. This

result indicates either that c-tubulin and microtubules
are not required for macronuclear division during veg-
etative growth in E. focardii or that the anti-a-tubulin
serum and the anti-c-tubulin sera used in the present
study are unable to recognize macronuclear a- and
c-tubulin isotypes in the E. focardii macronucleus,
perhaps due to post-translational modifications that
block the corresponding tubulin epitopes. We regard
the second explanation as being unlikely.
Cold adaptation of E. focardii c-tubulins –
implications for the mechanism of microtubule
nucleation
The suite of amino acid substitutions observed in
c
-T1 and c-T2 with respect to the Euplotes c-tubulin
(generally small polar residues replacing bulky and
charged residues) is similar to those that transform
mesophilic subtilisin-like proteases into cold-active
variants [54]. Similar trends in sequence substitutions
[55] have also been observed in the cold-adapted vari-
ants of lactate dehydrogenase [56], seralysin [57], the
pheromone En-1 from Euplotes nobilii [58], the ribo-
somal proteins P0 and P2 [32], two subunits of the
chaperonin containing TCP-1 [59], and a theromlysin-
like enzyme [60]. The overall effect of this substitu-
tion pattern is to introduce greater flexibility in the
cold-functioning proteins, particularly in surface loops
whose conformational adjustments are required to
facilitate attaining the active-site transition state in
cold thermal regimes.

How do the unique features of the E. focardii c-tubu-
lins enhance microtubule nucleation by the c-TuRC
complex at low temperature, and what do these changes
imply regarding the mechanism of nucleation? Based on
the evidence presented in the present study, we propose
that the conformational differences and increased
hydrophobicity of the M loops of c-T1 and c-T2 pro-
mote the lateral interactions necessary to form the
c-TuRC ring template at cold temperatures [12,35]
and that the residue substitutions and conformational
changes at the plus-end facilitate longitudinal inter-
action with the a-tubulin subunit located at the
minus-end of the tubulin heterodimer [12] or lateral
interactions with the b-tubulin subunit [61]. These
hypotheses are readily amenable to testing via site-direc-
ted mutagenesis and functional analysis of c-tubulins in
several model systems, including the yeasts Saccharo-
myces cerevisiae and Schizosaccharomyces pombe. Thus,
our comparative analysis of the c-tubulins of psychro-
philic and mesophilic Euplotes species should contribute
to resolving the validity of the template versus pro-
tofilament models of c-TuRC-mediated nucleation of
microtubules.
Experimental procedures
Cell culture and cell cycle synchronization
Cultures of E. focardii strains TN1 and TN15 were used
[27]; they represent type-species material chosen from a
number of wild-type strains isolated from sediment and
seawater samples collected in Antarctica (Terra Nova Bay
F. Marziale et al. c-Tubulin isotypes in E. focardii

FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5377
coastal waters). They were cultivated in a cold room
at 4 °C using the green alga Dunaliella tertiolecta as
food. Cells from logarithmic phase cultures were synchro-
nized by three consecutive treatments of starvation and
re-feeding.
Isolation of E. focardii c-tubulin
nanochromosomes from macronuclear
DNA via PCR and RATE
E. focardii macronuclear DNA was purified as described pre-
viously [31]. To obtain partial c-tubulin gene sequences, we
based our PCR strategy on degenerate oligonucleotide prim-
ers designed from a Euplotes c-tubulin consensus sequence
(obtained by the alignment of genes from three mesophilic
Euplotes; GenBank accession numbers: Euplotes aediculatus,
X85233.1; Euplotes otocarinatus, Y09553.1; E. crassus,
X85234.1 and X85235.1). The forward primer, 5¢-ATGCCA
AGAGAAATYATYACTTG-3¢, covered codons 1–7 plus
two nucleotides from 8, and the reverse primer, 5¢-TGAA
CTTGAGTTGGRTCAACRTC-3¢, corresponded to two
nucleotides of codon 328 plus triplets 327–321. Amplifica-
tion was performed using standard conditions: 30 cycles at
94 °C for 50 s, 45 °C for 1 min, and 72 °C for 1 min.
A final incubation at 72 °C for 7 min was added to the last
cycle. The amplification products were expected to contain
328 codons.
E. focardii macronuclear DNA is composed of small
nanochromosomes, each usually containing a single gene,
which are always terminated by telomeres consisting of four
repetitions of the motif C

4
A
4
[62]. This stereotypic organi-
zation facilitated obtaining the sequences of the C-terminal
coding region and the 5¢- and 3¢-UTRs using the RATE-
PCR technique as described previously [31,32]. We used the
forward and reverse primers (see above) individually in
combination with the telomeric oligonucleotide 5¢-(C
4
A
4
)
4
-
3¢. Amplified products were cloned into the pCR2.1-TOPO
vector of the TOPO TA Cloning
Ò
kit (Invitrogen, San
Diego, CA, USA) following the manufacturer’s recommen-
dations. Colony blotting and double-strand DNA labeling
by the random priming method were performed as
described previously [63]. Clones containing c-tubulin-
recombinant plasmids were sequenced in both strands (ABI
Prism sequence analyzer Model 373A and Big Dye Termi-
nator Methodology; PE Applied Biosystems, Foster City,
CA, USA).
DNA sequence analysis of c-tubulin
nanochromosomes and prediction of the
encoded amino acid sequences

DNA sequence analysis, amino acid sequence prediction,
and sequence alignments were performed with lasergene
Ò
,
version 7.2 (DNASTAR Inc., Madison, WI, USA).
Comparative protein structure modelling of E. focardii
c-tubulins
Comparative homology models of the two E. focardii c-tub-
ulins were obtained by use of modeller (version 9.1) [64]
and the friend interface [65]. The 3.0 A
˚
structure of
human c-tubulin containing bound GTP (Protein Databank
1z5w) [66] was used as a template for comparative mod-
elling. Structural alignments between the template and
modeled sequences were performed with topofit [66] and
models were analyzed under friend. The percentage simi-
larities between modeled and template sequences were
68.36% for c-T1 and 69.28% for c-T2, and the length of
alignment was 433 residues for both models. Based on these
values, we estimate that the accuracies of the modeled
structures of c-T1 and c-T2 approach 3 A
˚
[62].
Poly[A] + RNA purification, cDNA synthesis and
cloning, and Southern and northern blotting
Poly[A] + RNA was purified from E. focardii using the
Quick-Prep
Ò
mRNA purification kit (GE Healthcare Life

Sciences, Milan, Italy). For cDNA synthesis, the poly[A] +
RNA (4 lg) was treated with 10 U of RNase-free DNaseI
(Bethesda Research Laboratories, Bethesda, MD, USA), in
the presence of 40 U of RiboLock
Ò
(Fermentas, Milan,
Italy) and 4 mm MgCl
2
for 1 h at 37 °C. DNase-treated
RNA was incubated with Moloney murine leukemia virus
reverse transcriptase (Bethesda Research Laboratories) as
recommended by the manufacturer. The resulting cDNA
was then precipitated in ethanol, collected by centrifuga-
tion, resuspended in distilled water, and used as template
for PCR; the program was 30 cycles at 94 °C for 50 s,
48 °C for 1 min, and 72 °C for 1 min. A final incubation at
72 °C for 7 min was added to the last cycle. cDNA was
cloned into pCR2.1-TOPO as described above.
Southern and northern blotting were performed accord-
ing to standard procedures on Hybond-N filters from
Amersham (Milan, Italy). Filters were prehybridized,
hybridized to DNA probes, and washed to remove nonspe-
cifically bound probe according to the manufacturer’s rec-
ommendations. Filters were stripped for reuse by boiling in
distilled water for 15 s.
Estimation of macronuclear gene copy number and
gene transcription
To estimate gene copy number, qPCR was performed on
total macronuclear DNA using the SYBR green DNA-
binding method (TaKaRa Biotech, Dalian, China). The

two E. focardii c-tubulin genes were distinguished by use of
the following primer pairs: c-T1, EfgT1_FW (5¢-ATGCGT
CGTTTATTGCAGACT-3¢) as forward primer and Efg-
T1_REV (5¢-TGTTTTAGATAGAGCAACTTGGATT-3¢)
c-Tubulin isotypes in E. focardii F. Marziale et al.
5378 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
as reverse primer; and c-T2, EfgT2_FW (5¢-ATGCGTCGT
TTATTGCAACCC-3¢) as forward primer and Efg-T2_
REV (5¢-TGTTTCCGCTATCGATGTATGGTGA-3¢)as
reverse primer. 12.5 lL of SYBR Premix Ex Taq (2·) buf-
fer, 5 pg of each primer, and water were added to 100 ng
of E. focardii macronuclear DNA to reach a final volume
of 25 lL. The PCR parameters were initial denaturation at
95 °C for 2 min to activate the polymerase followed by 45
cycles of denaturation at 95 °C for 30 s and annealing and
extension at 60 °C for 15 s each. Following amplification,
melting curve analysis of the DNA was performed at tem-
peratures in the range 50–95 °C, with the temperature
increasing at a rate of 0.5 °C every 10 s. All PCR reactions
were performed in a Multicolor qPCR Detection System
iCycleriQ (Bio-Rad, Milan, Italy). During the primer
annealing ⁄ extension step, the increase in the fluorescence
from the amplified DNA was recorded by using the SYBR
Green optical channel set at a wavelength of 495 nm. The
initial threshold value was set at 30 fluorescent units. To
analyse transcription of the two c-tubulin mRNAs, the
same qPCR protocol was performed using 100 ng of
E. focardii cDNA prepared from total RNA.
Preparation of antibodies to E. focardii c-tubulins
Peptides that distinguish c-T1 [(390)RIFRRRNAYID-

NYK(403)] and c-T2 [(390)KKLRSNNAFITTYQ(403)]
were synthesized by Sigma Genosys (Milan, Italy) and used
as antigens to obtain rabbit polyclonal anti-peptide sera.
The c-T1 peptide was immunogenic, whereas the c-T2
peptide was not.
Immunofluorescence microscopy
E. focardii cells in logarithmic phase were washed, placed
on a polylysine-coated (0.5 mgÆmL
)1
) coverslip, and per-
meabilized with 0.2% Triton X-100 in PHEM buffer
(60 mm Pipes, 25 mm Hepes, 10 mm EGTA, 2 mm MgCl
2
,
final pH adjusted to 6.9 with NaOH). Cells were fixed with
2% paraformaldehyde in PHEM for approximately 30–
60 min, washed once with NaCl ⁄ P
i
(130 mm NaCl, 2 mm
KCl, 8 mm Na
2
HPO
4
,2mm KH
2
PO
4
, pH 7.2), and then
twice with NaCl ⁄ P
i

plus 0.1% BSA; washes were 10 min
each. Cells were incubated with rabbit polyclonal anti-
(human c-tubulin) serum (1 : 100 dilution in NaCl ⁄ P
i
) and
mouse monoclonal anti-a-tubulin serum DM1A (1 : 50;
Sigma Genosys) overnight at 4 °C, washed, and then incu-
bated with fluorescent secondary antibodies [Alexa Fluor
594 goat anti-(rabbit IgG) and Alexa Flour 488 goat anti-
(mouse IgG)] at 1 : 200 dilutions for 1 h at 37 °C; in some
experiments, rabbit polyclonal anti-(E. focardii c-T1) was
substituted for the anti-(human c-tubulin) serum. Finally,
cells were washed and suspended in 0.5% propyl gallate in
glycerol. Images were collected using an MRC600 Bio-Rad
confocal system connected to a Nikon inverted microscope
(Diaphot-TMD equipped with an Apoplan ·60 objective;
Nikon, Tokyo, Japan).
Preparation of whole cell extract, basal bodies, and
nuclei
Whole cell extract preparations were obtained as described
previously [29,31]. Basal bodies were extracted by the fol-
lowing procedure: (a) E. focardii cells were washed with
MT buffer [30 mm Tris–acetate (pH 7.3), 5 mm MgSO
4
,
5mm EGTA, 25 mm KCl, 1 mm dithiothreitol], collected
by low-speed centrifugation, and resuspended by extensive
stirring in two volumes of MT buffer containing 2% NP-40
and protease inhibitors (0.01% aprotinin, 0.005% phen-
ylmethanesulfonyl fluoride). An equal volume of 50%

(w ⁄ v) Percoll was then added, and the mixture was centri-
fuged for 30 min at 14 500 g in a fixed-angle rotor. The
fraction containing the basal bodies was recovered from the
interface between the Percoll and aqueous phases, diluted
in MT buffer, and centrifuged for 15 min at 14 500 g. The
resulting pellet was washed twice in MT buffer.
Nuclei were prepared by resuspension of E. focardii cell
pellets in two volumes of lysis buffer [10 mm Tris–HCl
(pH 6.8), 0.25 m sucrose, 10 mm MgCl
2
,1mm phen-
ylmethanesulfonyl fluoride, 0.5% NP-40] with gentle stirring
on ice for 2 min; two volumes of washing buffer (0.25 m
sucrose, 10 mm MgCl
2
) were added to stop lysis. The nuclear
suspension was centrifuged at 1000 g for 1 min (4 °C), the
supernatant was transferred to glass tubes, and nuclei were
collected by centrifugation (9000 g, 2 min, 4 °C).
SDS ⁄ PAGE and immunoblotting
Denaturing SDS ⁄ PAGE was performed according to the
method of Laemmli [67]. After electrophoresis, gels were
subjected to immunoblotting as described previously [29].
Blots were incubated either with a rabbit polyclonal anti-
(human c-tubulin) primary serum [40] at a 1 : 1000 dilution
or with a rabbit polyclonal antibody directed against the
c-T1 peptide (see above) at the same dilution. Blots were
washed extensively in NaCl ⁄ Tris ⁄ Tween buffer [5 mm Tris–
HCl (pH 7.5), 0.138 m NaCl, 0.1% Tween-20] and then
incubated with peroxidase-conjugated secondary anti-rabbit

serum (1 : 1000 dilution). After washing with NaCl ⁄
Tris ⁄ Tween, bound primary antibody was detected by
enhanced chemiluminescence (ECL Western Blotting
Analysis System; GE Healthcare).
Chemicals, materials, and reagents
DNA modifying and restriction enzymes, RNAse A,
32
P-dATP, and Hybond-N filters were purchased from
GE Healthcare. Taq polymerase was from PE Applied
Biosystems. Oligonucleotides were synthesized by Labtek
F. Marziale et al. c-Tubulin isotypes in E. focardii
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5379
Eurobio (Milan, Italy). All routine chemicals were of ana-
lytical grade and supplied by Sigma Aldrich (Milan, Italy).
Acknowledgements
We are grateful to Professor Piero Luporini for pro-
viding conceptual and practical advice during the
study, and Robin Leguy for anti-(human c-tubulin)
serum preparation. This research was supported by
grants from the Italian PNRA and MIUR to C.M., by
US National Science Foundation grant ANT-0635470
to H.W.D, and by US National Institutes of Health
grant R01LM009519 to V.A.I.
References
1 Dutcher SK (2003) Elucidation of basal body and cen-
trioles functions in Chlamydomonas reinhardtii. Traffic
4, 443–451.
2 Dawe HR, Farr H & Gull K (2007) Centriole ⁄ basal
body morphogenesis and migration during ciliogenesis
in animal cells. J Cell Sci 120, 7–15.

3 Oakley CE & Oakley BR (1989) Identification of
gamma-tubulin, a new member of the tubulin superfam-
ily by mipA gene of Aspergillus nidulans. Nature 338,
662–664.
4 Oakley BR, Oakley CE, Yoon Y & Jung MK (1990)
Gamma-tubulin is a component of the Spindle Pole
Body that is essential for microtubule function in Asper-
gillus nidulans. Cell 61, 1289–1301.
5 Sunkel CE, Gomes R, Sampaio P, Perdigao J & Gonz-
alez C (1995) Gamma-tubulin is required for the struc-
ture and function of the microtubule organizing centre
in Drosophila neuroblasts. EMBO J 14, 28–36.
6 Dammermann A, Mu
¨
ller-Reichert T, Pelletier L, Haber-
man B, Desai A & Oegema K (2004) Centriole assembly
requires both centriolar and pericentriolar material pro-
teins. Dev Cell 7, 815–829.
7 Ruiz F, Beisson J, Rossier J & Dupuis-Williams P
(1999) Basal body duplication in Paramecium requires
c-tubulin. Curr Biol 9, 43–46.
8 Shang Y, Li B & Gorovsky MA (2002) Tetrahymena
thermophila contains a conventional c-tubulin that is
differentially required for the maintenance of different
microtubule-organizing centers. J Cell Biol 158, 1195–
1206.
9 Moritz M & Agard DA (2001) c-Tubulin complexes
and microtubule nucleation. Curr Opin Struct Biol 11,
174–181.
10 Knop M, Pereira G, Geissler S, Grein K & Schiebel E

(1997) The spindle pole body component Spc97p inter-
acts with the gamma-tubulin of Saccharomyces cerevisi-
ae and functions in microtubule organizing and spindle
body duplication. EMBO J 16, 1550–1564.
11 Oegema K, Wiese C, Martin OC, Milligan RA, Iwama-
tsu A, Mitchison TJ & Zheng Y (1999) Characterization
of two related Drosophila gamma tubulin complexes
that differ in the ability to nucleate microtubules. J Cell
Biol 144, 721–733.
12 Kollman JM, Zelter A, Muller EG, Fox B, Rice LM,
Davis TN & Agard DA. (2008) The structure of the
{gamma}-tubulin small complex: implications of its
architecture and flexibility for microtubule nucleation.
Mol Biol Cell 19, 207–215.
13 Erickson HP & Stoffler D (1996) Protofilaments and
rings, two conformations of the tubulin family con-
served from bacterial FtsZ to alpha ⁄
beta and gamma
tubulin. J Cell Biol 135, 5–8.
14 Zheng Y, Wang ML, Alberts B & Mitchinson T (1995)
Nucleation of microtubule assembly by a c-tubulin-con-
taining ring complex. Nature 378, 578–583.
15 Job D, Valiron O & Oakley B (2003) Microtubule
nucleation. Curr Opin Cell Biol 15, 111–117.
16 Detrich HW III, Johnson KA & Marchese-Ragona SP
(1989) Polymerization of Antarctic fish tubulins at low
temperatures: energetic aspects. Biochemistry 28, 10085–
10093.
17 Detrich HW III, Fitzgerald TJ, Dinsmore JH & Mar-
chese-Ragona SP (1992) Brain and egg tubulins from

Antarctic fishes are functionally and structurally dis-
tinct. J Biol Chem 267, 18766–18775.
18 Herzog W & Weber K (1977) In vitro assembly of pure
tubulin into microtubules in the absence of microtu-
bule-associated proteins and glycerol. Proc Natl Acad
Sci USA 74, 1860–1864.
19 Himes RH, Burton PR & Gaito JM (1977) Dimethyl
sulfoxide-induced self-assembly of tubulin lacking asso-
ciated proteins. J Biol Chem 252, 6222–6228.
20 Melki R, Carlier M-F, Pantaloni D & Timasheff SN
(1989) Cold depolymerization of microtubules to double
rings: geometric stabilization of assemblies. Biochemistry
28, 9143–9152.
21 Billger M, Wallin M, Williams RC Jr & Detrich HW
III (1994) Dynamic instability of microtubules from
cold-living fishes. Cell Motil Cytoskel 28, 327–332.
22 Detrich HW III, Parker SK, Williams RC, Nogales E &
Downing KH (2000) Cold adaptation of microtubule
assembly and dynamics: structural interpretation of pri-
mary Sequence changes present in the a- and b-tubulins
of Antarctic fishes. J Biol Chem 275, 37038–37048.
23 Paluh JL, Killilea AN, Detrich HW III & Downing KH
(2004) Meiosis-specific failure of cell cycle progression
in fission yeast by mutation of a conserved b-tubulin
residue. Mol Biol Cell 15, 1160–1171.
24 Redeker V, Frankfurter A, Parker SK, Rossier J &
Detrich HW III (2004) Posttranslational modification of
brain tubulins from the Antarctic fish Notothenia
coriiceps: reduced C-terminal glutamylation correlates
c-Tubulin isotypes in E. focardii F. Marziale et al.

5380 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS
with efficient microtubule assembly at low temperature.
Biochemistry 43, 12265–12274.
25 Shang Y, Tsao CC & Gorovsky MA (2006) Mutational
analyses reveal a novel function of the nucleotide-bind-
ing domain of gamma-tubulin in the regulation of basal
body biogenesis. J Cell Biol 171, 1035–1044.
26 Jung MK, Prigozhina N, Oakley CE, Nogales E &
Oakley BR (2001) Alanine-scanning mutagenesis of
Aspergillus nidulans c-tubulin yields diverse and novel
phenotypes. Mol Biol Cell 12, 2119–2136.
27 Valbonesi A & Luporini P (1993) Biology of Euplotes
focardii an Antarctic ciliate. Polar Biol 13 , 489–493.
28 La Terza A, Papa G, Miceli C & Luporini P (2001)
Divergence between two Antarctic species of the ciliate
Euplotes, E. focardii and E. nobilii, in the expression of
heat-shock protein 70 genes. Mol Ecol 10, 1061–1067.
29 Pucciarelli S, Ballarini P & Miceli C (1997) Cold-
adapted microtubules: characterization of tubulin post-
translational modifications in the Antarctic ciliate Eupl-
otes focardii. Cell Motil Cytoskel 38, 329–341.
30 Miceli C, Pucciarelli S, Ballerini P & Luporini P (2000).
Cold-stable microtubules of the Antarctic ciliate Eupl-
otes focardii. In: Antarctic Ecosystems: Models for
Wider Ecological Understanding (Davison W, Howard-
Williams C & Broady P, eds), pp. 154–157. The Caxton
Press, Canterbury University, Christchurch.
31 Pucciarelli S & Miceli C (2002) Characterization of the
cold-adapted alpha-tubulin from the psychrophilic cili-
ate Euplotes focardii. Extremophiles 5, 385–389.

32 Pucciarelli S, Marziale F, Di Giuseppe G, Barchetta S
& Miceli C (2005) Ribosomal cold-adaptation: charac-
terization of the genes encoding the acidic ribosomal P0
and P2 proteins from the Antarctic ciliate Euplotes
focardii. Gene 7, 103–110.
33 Meyer F, Schmidt HJ, Plumper E, Hasilik A,
Mersmann G, Meyer HE, Engstrom A & Heckmann K
(1991) UGA is translated as cysteine in pheromone 3 of
Euplotes octocarinatus. Proc Natl Acad Sci USA 88,
3758–3761.
34 Tan M & Heckmann K (1998) The two c-tubulin-
encoding gene of the ciliate Euplotes crassus differ in
their sequences, codon usage, transcription initia-
tion sites and poly (A) addition sites. Gene 210, 53–60.
35 Aldaz H, Rice LM, Stearns T & Agard DA (2005)
Insights into microtubule nucleation from the crystal
structure of human gamma-tubulin. Nature 435, 523–
527.
36 Incla
`
n YF & Nogales E (2001) Structural models for
the self-assembly and microtubule interactions of
gamma- delta- and epsilon-tubulin. J Cell Sci 114, 413–
422.
37 Nogales E, Wolf SG & Downing KH (1998) Structure
of the alpha beta tubulin dimer by electron crystallogra-
phy. Nature 391, 199–203.
38 La Terza A, Miceli C & Luporini P (1995) Differential
amplification of pheromone genes of the ciliate Euplotes
raikovi. Dev Genet 17, 272–279.

39 Barchetta S, La Terza A, Ballarini P, Pucciarelli S &
Miceli C (2008) A combination of two regulatory ele-
ments in the Tetrahymena thermophila HSP70-1 gene
controls heat shock activation. Eukaryot Cell 7, 379–
386.
40 Fouquet JP, Kann ML, Combeau C & Melki R (1998)
Gamma-tubulin during the differentiation of spermato-
zoa in various mammals and man. Mol Hum Reprod 4,
1122–1129.
41 Go
¨
rtz HD (1982) The behavior and fine structure of the
dorsal bristles of Euplotes minuta, E. aediculatus, and
Stylonychia mytilus (Ciliata, Hypotrichida). J Eukaryot
Microbiol 29, 353–359.
42 Fleury A (1991) Dynamics of the cytoskeleton
during morphogenesis in the ciliate Euplotes. I. Basal
bodies related microtubular system. Eur J Protistol 27,
99–104.
43 Fleury A (1991) Dynamics of the cytoskeleton during
morphogenesis in the Ciliate Euplotes. II. Cortex and
continuous microtubular systems. Eur J Protistol 27,
220–237.
44 Liang A & Heckmann K (1993) The macronuclear
gamma-tubulin-encoding gene of E. octocarinatus
contains two introns and an in-frame TGA. Gene 22,
319–322.
45 Patient RK & McGhee JD (2002) The GATA family
(vertebrates and invertebrates). Curr Opin Gen Develop
12, 416–422.

46 Willem S, Srahna M, Devos N, Gerday C, Loppes R &
Matagne RF (1999) Protein adaptation to low tempera-
tures: a comparative study of alpha-tubulin sequences
in mesophilic and psychrophilic algae. Extremophiles 3,
221–226.
47 Paluh JL, Nogales E, Oakely BR, McDonald K, Pidoux
AL & Cande WZ (2000) A mutation in gamma-tubulin
alters microtubule dynamics and organization and is
synthetically lethal with the kinesin-like protein pkl1p.
Mol Biol Cell 11, 1225–1239.
48 Coleman DE, Berghuis AM, Lee E, Linder ME, Gil-
man AG & Sprang SR (1997) Structures of active con-
formations of Gi alpha 1 and the mechanism of GTP
hydrolysis. Science 265, 1405–1412.
49 Lambright DG, Sondek J, Bohm A, Skiba NP, Hamm
HE & Sigler PB (1996) The 2.0 A crystal structure of a
heterotrimeric G protein. Nature 379, 311–319.
50 Liang A, Ruiz F, Heckmann K, Klotz C, Tollon Y,
Beisson J & Wright M (1996) Gamma-tubulin is perma-
nently associated with basal bodies in ciliates. Eur J
Cell Biol 70
, 331–338.
51 Joachimiak E, Pucciarelli S, Barchetta S, Ballarini P,
Kaczanowska J & Miceli C (2006) Cell cycle-dependent
F. Marziale et al. c-Tubulin isotypes in E. focardii
FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS 5381
expression of gamma-tubulin in the amicronuclear cili-
ate Tetrahymena pyriformis. Protist 158, 39–50.
52 Klotz C, Ruiz F, Garreau de Loubresse N, Wright M,
Dupuis-Williams P & Beisson J (2003) Gamma-tubulin

and MTOCs in Paramecium. Protist 154, 193–209.
53 Fujiu K & Numata O (2000) Reorganization of micro-
tubules in the amitotically dividing macronucleus of
Tetrahymena. Cell Motil Cytoskeleton 46, 17–27.
54 Wintrode PL, Miyazaki K & Arnold FH (2000)
Cold adaptation of a mesophilic subtilisin-like prote-
ase by laboratory evolution. J Biol Chem 275, 31635–
31640.
55 Borders CL Jr, Broadwater JA, Bekeny PA, Salmon
JE, Lee AS, Eldridge AM & Pett VB (1994) A struc-
tural role for arginine in proteins: multiple hydrogen
bonds to backbone carbonyl oxygens. Protein Sci 4,
541–548.
56 Fields PA & Somero GN (1998) Hot spots in cold
adaptation: localized increases in conformational flexi-
bility in lactate dehydrogenase A4 orthologs of Antarc-
tic notothenioid fishes. Proc Natl Acad Sci USA 95,
11476–11481.
57 Kumeta H, Hoshino T, Goda T, Okayama T,
Shimada T, Ohgiya S, Matsuyama H & Ishizaki K.
(1999) Identification of a member of the serralysin
family isolated from a psychrotrophic bacterium,
Pseudomonas fluorescens 114. Biosci Biotechnol
Biochem 63, 1165–1170.
58 Alimenti C, Ortenzi C, Carratore V & Luporini P
(2003) Structural characterization of En-1, a cold-
adapted protein pheromone isolated from the Antarctic
ciliate Euplotes nobilii . Biochimica and Biophysica Acta
1621, 17–21.
59 Pucciarelli S, Parker SK, Detrich HW III & Melki R.

(2006) Characterization of the cytoplasmic chaperonin
containing TCP-1 from the Antarctic fish Notothenia
coriiceps. Extremophiles 10, 537–549.
60 Adekoya OA, Helland R, Willassen NP & Sylte I
(2006) Comparative sequence and structure analysis
reveal features of cold adaptation of an enzyme in the
thermolysin family. Proteins: Structure, Function, and
Bioinformatics 62, 435–449.
61 Leguy R, Melki R, Pantaloni D & Carlier MF (2000)
Monomeric gamma-tubulin nucleates microtubules. J Biol
Chem 275, 21975–21980.
62 Hoffman DC, Anderson RC, DuBois ML & Prescott
DM (1995) Macronuclear gene-sized molecules of hypo-
trichs. Nucleic Acids Res 23, 1279–1283.
63 Sambrook J & Russel DW (2001) Molecular Cloning:
A Laboratory Manual, 3rd edn. Cold Spring Harbor
Laboratory Press, New York, NY.
64 Martı
´
-Renom MA, Stuart AC, Fiser A, Sanchez R,
Melo F & Sali A (2000) Comparative protein structure
modelling of genes and genomes. Annu Rev Biophys
Biomol Struct 29, 291–325.
65 Abyzov A, Errami M, Leslin CM & Ilyin VA (2005)
Friend, an integrated analytical front-end application
for bioinformatics. Bioinformatics 21, 3677–3678.
66 Ilyin VA, Abyzov A & Leslin CM (2004) Structural
alignment of proteins by a novel TOPOFIT method, as
a superimposition of common volumes at a topomax
point. Protein Sci 13

, 1865–1874.
67 Laemmli UK (1970) Cleavage of structural proteins
during the assembly of the head of bacteriophage T4.
Nature 227, 680–685.
68 Llanos R, Chevrier V, Ronjat M, Meurer-Grob P,
Martinez P, Frank R, Bomens M, Wade RH, Wehland
J & Job D (1999) Tubulin binding sites on gamma-
tubulin identification and molecular characterization.
Biochemistry 38, 15712–15720.
69 Pettersen EF, Goddard TD, Huang CC, Couch GS,
Greeenblatt DM, Meng EC & Ferrin TE (2004) UCSF
Chimera – a visualization system for exploratory
research and analysis. J Comput Chem 25, 1605–1612.
Supporting information
The following supplementary material is available:
Fig. S1. Comparative modelling of E. focardii c-tubu-
lins using the structure of human c-tubulin as a
template.
This supplementary material can be found in the
online version of this article.
Please note: Wiley-Blackwell is not responsible for
the content or functionality of any supplementary
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than missing material) should be directed to the
corresponding author for the article.
c-Tubulin isotypes in E. focardii F. Marziale et al.
5382 FEBS Journal 275 (2008) 5367–5382 ª 2008 The Authors Journal compilation ª 2008 FEBS