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Cell Motility and the Cytoskeleton 39:318–330 (1998)
Tubulin Polyglycylation in Platyhelminthes:
Diversity Among Stable Microtubule
Networks and Very Late Occurrence
During Spermiogenesis
Carlo Iomini,1 Marie-Hélène Bré,2 Nicolette Levilliers,2 and Jean-Lou Justine1*
1Laboratoire
de Biologie Parasitaire, Protistologie, Helminthologie, ERS 156
CNRS, Muséum National d’Histoire Naturelle, Paris, France
2Laboratoire de Biologie Cellulaire 4, URA 2227 CNRS, Université de Paris-Sud,
Orsay, France
The distribution of glycylated tubulin has been analyzed in different populations of
stable microtubules in a digenean flatworm, Echinostoma caproni (Platyhelminthes). Two cellular types, spermatozoa and ciliated excretory cells, have been
analyzed by means of immunofluorescence, immunogold, and immunoblotting
techniques using two monoclonal antibodies (mAbs), AXO 49, and TAP 952,
specifically directed against differently glycylated isoforms of tubulin. The
presence of glycylated tubulin in the two cell types was shown. However, the
differential reactivities of TAP 952 and AXO 49 mAbs with the two axoneme types
suggest a difference in their glycylation level. In addition, within a single cell, the
spermatozoon, cortical microtubules underlying the flagellar membrane, and
axonemal microtubules were shown to comprise different tubulin isoforms, the
latter ones only being labelled with one of the antiglycylated tubulin mAbs, TAP
952. Similarly, the antiacetylated (6-11B-1) and polyglutamylated (GT335) tubulin
mAbs decorated the two types of axonemal microtubules, but not the cortical ones.
From these data, a subcellular sorting of posttranslationally modified tubulin
isoforms within spermatozoa, on the one hand, and a cellular sorting of glycylated
isoforms inside the whole organism, on the other hand, is demonstrated in the
flatworm E. caproni. Last, a sequential occurrence of tubulin posttranslational
modifications was observed in the course of spermiogenesis. Acetylation appears
first, followed shortly by glutamylation; glycylation takes place at the extreme end
of spermiogenesis and, specifically, in a proximo-distal process. Thus in agreement
with, and extending other studies [Bré et al., 1996], glycylation appears to close the
sequence of posttranslational events occurring in axonemal microtubules during spermiogenesis. Cell Motil. Cytoskeleton 39:318–330, 1998. r 1998 Wiley-Liss, Inc.
Key words: posttranslational modifications; polyglycylation; stable microtubules; spermatozoa; Platyhelminthes
INTRODUCTION
In most eukaryotic cells, the diversity of the various
microtubule subpopulations is increased not only by the
presence of a- and b- tubulin isotypes encoded by
different multigenic families, but also by posttranslational
modifications (PTMs) of primary gene products.
r 1998 Wiley-Liss, Inc.
*Correspondence to: Jean-Lou Justine, Laboratoire de Biologie Parasitaire, Protistologie, Helminthologie, Muséum National d’Histoire
Naturelle, 61 rue Buffon, F-75231 Paris cedex 05, France. E-mail:
justine@mnhn.fr
Received 16 December 1997; accepted 23 December 1997
Tubulin Polyglycylation in Stable Microtubules
These PTMs can be subdivided into two classes.
One class includes the addition/removal of (an) amino
acid(s), or the chemical modification of a residue of the
polypeptide chains. With the exception of acetylation
[L’Hernault and Rosenbaum, 1985; LeDizet and Piperno,
1987] and palmitoylation [Caron, 1997; Ozols and Caron,
1997], PTMs, such as detyrosination [Barra et al., 1974;
Thompson, 1982], excision of the penultimate glutamate
residue [Paturle-Lafanechère et al., 1991], and phosphorylation [Eipper, 1974; see Khan and Ludueña, 1996],
occur in the carboxy-terminal region of tubulin. The
second class, comprising the polymodifications, has been
characterized by the lateral branching of a polyglutamyl
[Eddé et al., 1990], or a polyglycyl [Redeker et al., 1994]
chain of variable length on a specific glutamate residue
near the carboxy-terminal end of the tubulin subunits. A
wide panel of isoforms is thus generated by these
polymodifications.
The role of the tubulin PTMs is poorly known;
nevertheless, a general but not universal correlation has
been established between the stability of microtubules
and the extent of modification [for reviews, see Greer and
Rosenbaum, 1989; Gelfand and Bershadsky, 1991]. Accordingly, the polyglutamyl chain length has been assumed to regulate interactions of tubulin with microtubuleassociated proteins (MAPs) [Boucher et al., 1994]. In
addition, the polymodifications have been hypothesized
to play a role in spermatozoon motility [Bré et al., 1996;
Gagnon et al., 1996].
Using specific antibodies, we have investigated the
occurrence of tubulin PTMs, and particularly polyglycylation, in Echinostoma caproni, a digenean flatworm belonging to the phylum Platyhelminthes. This biological model
allowed us to address the question of the distribution of
glycylated isoforms among stable microtubules differently located inside a single organism. The access to germ
and excretory cells has permitted analysis of two cell
types, either flagellated (spermatozoa) or ciliated (protonephridia), both comprising motile axonemes arranged in
different patterns (9 1 ‘‘1’’ and 9 1 2, respectively) [Iomini and Justine, 1997]. In addition, the co-existence in
the spermatozoon of E. caproni of axonemes and cortical
microtubules allowed us to approach the question of the
distribution of polyglycylation comparatively to some
other PTMs in two types of stable microtubules—cortical
and axonemal—closely located within a same cell.
We showed that polyglycylation, as well as acetylation and polyglutamylation, were indeed present, but
differently distributed in the three classes of microtubules. Moreover, two complementary tools, specifically
directed against distinct levels of glycylation, were found
to be differentially reactive with the 9 1 ‘‘1’’ and 9 1 2
axonemes. These results show that the PTMs allow
319
discrimination between different functional classes of
stable microtubules inside a single organism or cell.
Because in E. caproni the different stages of
spermatid maturation are readily distinguishable, we
could accurately follow the time occurrence of glycylation, comparatively to acetylation and glutamylation, in
the course of spermiogenesis. This analysis revealed the
sequential appearance of these PTMs, glycylation being
the last one to take place, and delayed to the end of
spermiogenesis. The results are discussed in regard to the
mechanism of sorting and appearance of tubulin isoforms
in a single or in two compartments of one organism.
MATERIALS AND METHODS
Animals, Tissue, and Cell Isolation
Echinostoma caproni, a parasitic species from the
intestine of a wild bird [Richard, 1964], has been adapted
to laboratory mice and is widely used as a model [Fried
and Huffman, 1996]. Specimens were collected from
mice experimentally infested with 50 metacercariae. The
worms were dissected in phosphate-buffered saline (PBS)
at pH 7.4 in order to isolate either the testes or the seminal
vesicle. Spermatozoa were extracted from the seminal
vesicle with needles. Testes or spermatozoa were collected in PBS containing protease inhibitors [Bré et al.,
1996].
Antibodies
The monoclonal antibodies, TAP 952 and AXO 49,
raised against Paramecium axonemal tubulin [Callen et
al., 1994], specifically recognize different levels of tubulin polyglycylation [Bré et al., 1996]. AXO 49 ascitic
fluid was prepared by Dr. Jeanmaire-Wolf. GT335 mAb,
directed against polyglutamylated tubulin [Wolff et al.,
1992], was a kind gift from Dr. P. Denoulet (Collège de
France, Paris). 6-11B-1 mAb, directed against acetylated
a-tubulin [Piperno and Fuller, 1985], was purchased from
Sigma (France). DM1A and DM1B anti-a- and b-tubulin
mAbs [Blose et al., 1984] were purchased from Amersham (France). C105 polyclonal antibody (Ab), directed
against a carboxy-terminal sequence of b-tubulin [Arevalo et al., 1990], was kindly provided by Dr. J. M.
Andreu (Centro de Investigationes Biologicas, Madrid,
Spain).
The FITC- and TRITC-conjugated goat antimouse
and goat antirabbit IgG Abs were purchased from Nordic
(France). The gold-conjugated goat antimouse IgG Ab
was purchased from BioCell (France). Goat immunoglobulins were purchased from Sigma. The peroxidase-labelled
goat antimouse IgG Ab was from Sanofi Diagnostics
Pasteur (France).
320
Iomini et al.
Immunofluorescence
Germ cells were obtained by squashing testes or
seminal vesicles in a drop of PBS on a pit slide. Slides
were dried for 1 h under a fan, then kept at 4°C and
processed within 24 h. The cells were treated with
acetone for 10 min at room temperature and rinsed in PBS
(3 3 5 min). After rinsing, nonspecific antigens were
blocked with 2% bovine serum albumin (Sigma) in PBS
(BSA-PBS) for 45–90 min at room temperature. The pits
were then covered for 40 min with one antibody diluted in
BSA-PBS: AXO 49 cell culture supernatant, 1:2, or
ascitic fluid, 1:100; TAP 952, 1:10; 6-11B-1, 1:200;
GT335, 1:500; DM1A, 1:200; C105, 1:50. After rinsing
in PBS (3 3 5 min), a FITC goat anti-mouse IgG Ab
(1:40 in PBS) was applied for 40 min at room temperature.
For double labelling, the pits were successively
covered for 40 min with one of the antitubulin PTM Abs
cited above, then with C105 Ab, 1:50, then with FITCconjugated goat antimouse, and TRITC-conjugated goat
antirabbit IgG Abs, both 1:40, followed each time by
rinsing in PBS (3 3 5 min). Thereafter, nucleus staining
was achieved using DAPI (1 µg/ml in PBS) for 10 min.
After final rinsing in PBS (3 3 5 min), mounting was
carried out in CITIFLUOR (Citifluor, UK), and slides
were sealed with nail enamel. Controls were realized by
omitting the primary Abs or by using nonrelevant mouse
secondary Abs (they were negative and are thus not
further mentioned or illustrated). Observations were
made with a Nikon Optiphot epifluorescence microscope.
Postembedding Electron
Microscope Immunocytochemistry
Living specimens cut into pieces were fixed for 1 h
at 4°C in 2% glutaraldehyde in 0.1 M sodium cacodylate
at pH 7.2. After rinsing in the same buffer (3 3 10 min),
the worms were dehydrated in a graded ethanol series and
embedded in LR White resin, medium grade (Pelanne
Instruments, Toulouse, France). The resin was polymerized in tightly capped gelatin capsules for 10 h at 60°C.
Ultrathin sections were placed on nickel or gold grids.
Grids were rinsed (PBS, 3 3 5 min), and nonspecific
antigens were blocked with goat Igs (1:30 in PBS) for 1 h.
The grids were then incubated in a moist chamber
for 40 min at room temperature with a primary antibody
(AXO 49 ascitic fluid, 1:100; TAP 952, 1:4; 6–11B-1,
1:50; GT335, 1:500; DM1A, 1:100), diluted in PBS with
goat Igs (1:100). After rinsing in PBS (3 3 5 min), a
gold-conjugated goat antimouse IgG Ab (15 nm gold
beads, 1:20 in PBS) was applied for 1 h at room
temperature. After final rinsing in PBS (3 3 5 min,
followed by distilled water, 3 3 5 min), the grids were
dried on Whatman paper, stained according to Daddow
[1986], and observed with a Hitachi H600 electron
microscope.
Gel Electrophoresis and Immunoblotting
A whole worm was cut and crushed with a pestle in
a plastic tube containing liquid nitrogen and immediately
boiled in Laemmli buffer for 4 min. Testes and spermatozoa were resuspended in the same buffer and similarly
processed; proteins were separated in 10% polyacrylamide minigel-gels [Laemmli, 1970], containing 0.1%
(w/v) SDS, 99% pure (BDH, U.K.) or 70% pure (Sigma),
respectively, adjusted to pH 8.8 or 9.3. After electrophoresis, gels were stained with Coomassie brilliant blue R250
or electro-transferred onto nitrocellulose according to
Kyhse-Andersen [1984]. The blots were stained with
Ponceau red, washed in PBS, saturated with PBS containing 3% BSA and 0.1% Tween-20, and then incubated
with TAP 952, 1:50, AXO 49 ascitic fluid, 1:5,000,
GT335, 1:50,000, 6-11B-1, 1:30,000, DM1A, 1:5,000 or
DM1B, 1:1,000, diluted in PBS containing 0.3% BSA
and 0,1% Tween-20. After extensive washings with the
latter buffer, blots were incubated with peroxidaselabelled goat antimouse IgG Ab, 1:2,000, and processed
for enhanced chemiluminescence detection (ECL, Amersham). Some blots were reprobed with another primary
antibody and similarly processed.
RESULTS
Distribution of Polyglycylated Tubulin in Various
Classes of Stable Microtubules in Platyhelminthes
Although tubulin was a minor polypeptide with
respect to the whole protein amount of Echinostoma
caproni (Fig. 1, lanes 1 and 2), TAP 952 exhibited a
specific reactivity with two polypeptide bands migrating
at the same level as the a- and b-tubulin subunits
revealed by DM1A and DM1B. In contrast, a very weak
staining was obtained with AXO 49 (Fig. 1), an observation that could mean that a- and b-tubulin subunits of E.
caproni are only glycylated at low levels. To check this
assumption, we have comparatively investigated two
distinct axonemal systems expected to be enriched in
polyglycylated tubulin, namely, flagella of mature spermatozoa and cilia of protonephridia.
The seminal vesicle and the two testes were recovered by dissection in order to separate the mature
spermatozoa from the spermiogenesis stages. The spermatozoon of E. caproni is a long and filiform motile cell, in
which ultrastructural studies have demonstrated the presence of two parallel axonemes and cortical longitudinal
microtubules. The axonemes display the typical 9 1 ‘‘1’’
pattern of parasitic Platyhelminthes, i.e., they are made
up of the usual nine microtubule doublets and of a central
core 600 nm in diameter [Iomini et al., 1995].
Tubulin Polyglycylation in Stable Microtubules
Fig. 1. Immunoblot analyzis of proteins from the flatworm E. caproni.
The proteins were submitted to SDS-PAGE and stained with Coomassie Blue (C B: lane 1, E. caproni proteins; lane 2, calf brain
tubulin) or transferred onto nitrocellulose and incubated with DM1A
(a-tubulin), DM1B (b-tubulin), TAP 952, and AXO 49 (glycylated
tubulin). In order to improve the separation between alpha- and
beta-tubulin, the migration time of proteins has been increased for
immunoblotting compared to C.B.
Double staining immunofluorescence (Fig. 2) as
well as immunogold labelling (Fig. 3) revealed that
spermatozoa were only reactive with TAP 952, but not
with AXO 49. Similar reactivity characteristics were
observed after immunoblotting of proteins from seminal
vesicle spermatozoa (Fig. 4).
Close inspection of the distribution of gold particles
in transverse sections of spermatozoa showed that the
TAP 952 labelling concerned only the peripheral doublets
of axonemes but not the cortical microtubules (Fig. 3; see
also Fig. 9). This allowed the interpretation of double
labelling immunofluorescence observations (Fig. 2). With
the C 105 antitubulin sequence Ab, the single heavy line
corresponds to the axonemes and the thin more straight
line to the cortical microtubules, separated from the
axonemes by a fixation artefact. In contrast, with TAP
952, the thin line, corresponding to the cortical microtubules, is not revealed.
In order to determine whether the specific PTM
distribution found in spermatozoan microtubules concerned glycylation only, the reactivity of mAbs directed
against other tubulin PTMs was also investigated by
immunofluorescence and immunoelectron microscopy.
6-11B-1 and GT335 mAbs unequivocally labelled the
peripheral doublets of the axonemes, but not the cortical
microtubules, although the latter were sometimes weakly
labelled with GT335 (Table I). It is noteworthy that, with
321
Fig. 2. Double labelling immunofluorescence of E. caproni spermatozoan microtubules with C105 (b-tubulin) (a, c) and TAP 952 (glycylated tubulin) (b) or AXO 49 (glycylated tubulin) (d). The arrow
shows the cortical microtubules. Bar 5 50 µm.
all antitubulin Abs tested at the ultrastructural level, the
central core of the axonemes was never labelled (Fig. 3;
see also Fig. 9). This observation strengthens the statement that it does not contain tubulin [Iomini et al., 1995],
thus conferring on these axonemes a structural and
molecular singularity.
We took advantage of the presence in E. caproni of
another cell type possessing the classical axonemal
structure to check whether the exclusive reactivity of TAP
952 compared to AXO 49 might be a characteristic
feature of the whole organism.
The excretory system in most Platyhelminthes is
constituted by protonephridia [Rohde, 1990]. These cells
comprise a variable number of motile cilia arranged
according to a 9 1 2 pattern. Postembedding immunocytochemistry experiments revealed a labelling of the
peripheral doublets and central microtubules of the
axonemes with both antibodies directed against glycylated tubulin, AXO 49 and TAP 952 (Fig. 3a, b, d; Table
I). This suggests that a wide panel of glycylated isoforms
is present in protonephridia, in contrast to spermatozoa.
The exact number of protonephridia in E. caproni is
unknown, but these cells represent a small percentage of
the total body mass that can account for the weak staining
observed after immunoblotting of total protein extracts
with AXO 49 (Fig. 1). Nevertheless, one cannot exclude
the possibility that some deglycylation might occur
during the extract preparation. Concerning the other
322
Iomini et al.
Fig. 3. Immunogold labelling of ultrathin sections of protonephridia (a, b, d, f) and spermatozoa (c, e, g)
of E. caproni with AXO 49 (glycylated tubulin) (a, b, c), TAP 952 (glycylated tubulin) (d, e), and DM1A
(a-tubulin) (f, g). (a) Longitudinal section of a protonephridium, showing its general morphology; inset,
higher magnification. (b–g), Transversal sections. Bar 5 2 µm for (a), and 250 nm for inset and (b–g).
PTMs, the ciliary axonemes, including the peripheral
doublets and the two central microtubules of protonephridia, were likewise decorated with 6-11B-1 and GT335
mAbs (Table I).
Occurrence of Glycylation in the Course
of Spermiogenesis
Figure 4 shows comparative immunoreactivities of
protein extracts of testes and of spermatozoa, originating
from the seminal vesicles, with the different antitubulin
PTM Abs. Since very different protein amounts were
present in both extracts (Fig. 4, lanes 1, 2), indicating
various numbers of germ cells in the two compartments,
their respective loads were adjusted using DM1A and
DM1B reactivities as markers of their tubulin content.
AXO 49 was unreactive with either tubulin. Whereas
6-11B-1 and GT335 reactivities were approximately
similar for testis and seminal vesicle tubulin, TAP 952
exhibited a higher reactivity with the latter tubulin,
suggesting that glycylation takes place progressively
during spermiogenesis in the testes.
Ultrastructural studies [Iomini and Justine, 1997]
have shown that spermiogenesis in E. caproni follows a
peculiar morphological pattern (diagrammed in Fig. 6).
Each spermatid has a conical structure from which three
cell processes grow and progressively fuse, a median
cytoplasmic one, containing cortical microtubules, and
two flagella. Using specific fluorescent markers of tubulin
and DNA, the length of these processes and the position
of the nucleus can be visualized, thus readily permitting
identification of the stages of spermiogenesis. As illustrated in Figure 5, groups of cells with round nuclei (Fig.
5b) and short axonemes (Fig. 5a) correspond to early
spermatids (Fig. 6a). The axonemes and the cortical
microtubules lengthen (Figs. 5c, 6b) and fuse with the
median process (Figs. 5e, 6c) so that two axonemes,
surrounded by cortical microtubules, become incorporated within the spermatozoon (Figs. 5i, 6e). Simultaneously, the nuclei lengthen and migrate into the elongating spermatids (Figs. 5e, f, 6c), to be located eventually at
the distal extremity of the late spermatids (Figs. 5g, h, 6d)
and of the spermatozoa (Figs. 5i, j, 6e).
In order to specify the time occurrence of glycylation during spermiogenesis, spermatids from the testes
were analyzed by immunofluorescence with the antitubulin PTM antibodies (Fig. 7).
Tubulin Polyglycylation in Stable Microtubules
323
ylated isoforms appear first in the proximal part of the
axonemes and thereafter along the whole flagella to yield
mature spermatozoon.
As expected from the immunocytochemistry and
immunoblotting data, none of the spermiogenesis stages
within the testes was detected by AXO 49.
DISCUSSION
Fig. 4. Comparison of the extent of tubulin PTMs in protein extracts
from testes and from seminal vesicle (SV) spermatozoa of E. caproni.
For Coomassie Blue (C B), loads correspond to 0.7 testis (lane 1) or 0.7
vesicle (lane 2); for immunoblotting with DM1A (a-tubulin), DM1B
(b-tubulin), 6-11B-1 (acetylated tubulin), GT335 (glutamylated tubulin), TAP 952 and AXO 49 (glycylated tubulin), loads correspond to 0.3
testis or 3 vesicles. The migration of calf brain tubulin is indicated in
lane 3.
6-11B-1 mAb (Fig. 7b) revealed the early stages
(Fig. 7a, c), and all other stages throughout. GT335 (Fig.
7e) also detected the early stages (Fig. 7d, f), but at that
time, the decoration was restricted to the proximal part of
the elongating spermatids (compare Fig. 7d, 7e); the
whole length of the spermatids became labelled later (Fig.
7h), when the axonemes were elongating (Fig. 7g) and the
nuclei migrating (Fig. 7i). The labelling intensity seemed
to increase with the progression of spermiogenesis (Fig.
8), yielding fully labelled (Fig. 7k, n) late spermatids
(Fig. 7j, l) and spermatozoa (Fig. 7m, o). The TAP 952
epitope (Fig. 7q) was not detected until the last stages of
spermiogenesis, namely, when nuclear migration was
completed (Fig. 7p, r); only the proximal part of the
flagella was visible at this stage (compare p and q, Fig. 7).
The whole length of the flagella was labelled in mature
spermatozoa (Fig. 7t), once they were released from the
common mass (Fig. 7s). The differential distribution of
the TAP 952 epitope along the length of late spermatids
was checked by immunoelectron microscopy (Fig. 9). In
contrast to GT335, which stained the different parts of
late spermatids (Fig. 9a–c), only sections of such spermatids devoid of nucleus, interpreted as proximal, were
labelled with TAP 952 (Fig. 9d), but sections from the
middle part of spermatids (Fig. 9e), as well as distal ones
(Fig. 9f), were not labelled. This confirmed that glyc-
A series of observations carried out with specific
antibodies has allowed to infer that glycylation is widely
spread among axonemal tubulins, from ciliated protozoa
to Metazoa [Adoutte et al., 1991; Bressac et al., 1995;
Levilliers et al., 1995], and is lacking in neuronal tubulin
[Callen et al., 1994], in contrast to acetylation and
glutamylation, which have been found in both tubulin
types [see LeDizet and Piperno, 1991; for a review on
acetylation, see Wolff et al., 1992; Bré et al., 1994;
Fouquet et al., 1994]. The immunological tools have also
permitted recognition of different levels of glutamylation
in the mammalian cells [Audebert et al., 1993; Fouquet et
al., 1994] and of glycylation in phylogenetically distant
species ranging from ciliates to primates [Bré et al.,
1996]. In parallel, mass spectrometry analyses have
established that the number of glutamyl and of glycyl
units added to the tubulin subunits varies considerably
according to the organism analyzed [Eddé et al., 1990;
Alexander et al., 1991; Redeker et al., 1992, 1994; Rüdiger
et al., 1992, 1995; Mary et al., 1994, 1996; Multigner et
al., 1996; Weber et al., 1996; Schneider et al., 1997].
This study, revealing the presence of both polymodifications in E. caproni, extends knowledge of their
occurrence to the phylum Platyhelminthes. Both a- and
b-tubulin subunits have been found to be modified. The
distribution of these PTMs, and particularly glycylation,
has been analyzed in different populations of stable
microtubules in two cellular types of E. caproni, and their
time occurrence has been followed during spermiogenesis.
Three major results have been obtained: (1) in the
spermatozoon, two distinct functional classes of stable
microtubules comprise different tubulin isoforms, (2) in
the same organism, tubulin constituting two types of
axonemes has been shown, for the first time, to possess
different levels of glycylation, and (3) tubulin glycylation
takes place only at the final step of spermiogenesis, in
contrast to the early occurrence of acetylation and
glutamylation.
PTMs Discriminate Between Two Different
Functional Classes of Stable Microtubules
Within a Single Cell
Numerous studies have shown a correlation between the stability of microtubules and the presence of
324
Iomini et al.
TABLE I. Labelling of Microtubules With Antibodies Directed Against Various Tubulin PTMs
Cortical microtubules
spermatozoa
Axonemal microtubules
spermatozoa
protonephridia
AXO 49
(glycylation)
TAP 952
(glycylation)
6-11B-1
(acetylation)
GT335
(glutamylation)
2
2
2
1/2
2
1
1
1
1
1
1
1
*1 labelling, 2 no labelling, 1/2 weak labelling sometimes observed.
Fig. 5. Immunofluorescence staining of spermatid microtubules with
DM1A (a-tubulin) (a, c, e, g, i), and DNA labelling (b, d, f, h, j)
showing the time course of spermiogenesis in E. caproni. (a, b) Early
spermatids; (c, d) elongating spermatids with nuclei still in the central
mass; (e, f) distal part of elongating spermatids, showing the two
axonemes (arrowheads), partially fused (arrow), and migrating nuclei;
(g, h) late spermatids, with completely fused axonemes and distal
nuclei; i, j: mature spermatozoon with distal nucleus. Bar 5 50 µm.
Tubulin Polyglycylation in Stable Microtubules
Fig. 6. Diagram of morphological pattern of spermiogenesis in
E. caproni (adapted from Justine, 1991). Early spermatids (a)
show three elongating processes, a median cytoplasmic process (M)
and two flagella (F). The two flagella fuse with the median cytoplasmic
process in a proximo-distal pattern (b, c) and, simultaneously, the
nucleus (n) migrates into the distal extremity of the elongating
spermatid (b, c, d). Finally, the spermatozoon (e), containing two
incorporated axonemes, separates from the spermatid mass. Straight
arrows indicate elongation, curved arrows indicate the movement of
fusion of the flagella. Possible coiling of nucleus in the distal extremity
is not pictured.
PTMs, the latter thus reflecting a low microtubule turnover. However, it is worth comparing the distribution of
PTMs among the most stable microtubule classes of a
same cell in order to determine whether some differentiation can still be found among them. In mammalian
spermatids, a differential PTM distribution has in fact
been observed between two microtubule arrays, a transient one, that of the manchette, and a permanent one, that
of the flagellar axoneme [Fouquet et al., 1994]. The
simultaneous presence of cortical and axonemal microtubules in ciliated and flagellated cells provides an opportunity to analyze the PTM distribution among permanent
stable microtubules.
In the ciliate Paramecium, the three PTMs, acetylation, glutamylation, and glycylation, are located both in
the ciliary axonemes and in most cortical microtubules
[Bré et al., 1994; Fleury et al., 1995]. In contrast, in E.
caproni spermatozoa, comprising two 9 1 ‘‘1’’ axonemes
and cortical microtubules, the presence of the three PTMs
is restricted to the axonemal doublets, excluding the
neighbouring cortical microtubule singlets. Since both
325
microtubular systems are long-lived, their differential
modifications cannot be explained by different microtubule dynamics. A masking effect of the three PTM
epitopes by MAPs is also unlikely, given the distinct
locations of these PTMs on the tubulin molecule. Rather,
one mechanism could set in action a subcellular sorting of
tubulin isotypes toward the two microtubular networks
involved in different functions, as postulated in the
multitubulin hypothesis [Fulton and Simpson, 1976]. As a
matter of fact, a few cases of segregation and selective
utilization of isotypes have been reported for the construction of axonemes. For example, in the Drosophila male
germ line, the postmitotic b2 isotype, although multifunctional, i.e., required for the assembly of cytoplasmic
microtubules as well as for the sperm axoneme doublet
microtubules, has been shown to be indispensable for the
right construction of the axoneme [see Raff, 1994]. In the
framework of this hypothesis, one would have to consider
an isotype-specific occurrence of PTMs, as demonstrated
for phosphorylation [Khan and Ludueña, 1996]. As the
extreme carboxy-terminal domain is the most divergent
part of the tubulin molecule, one could assume, in E.
caproni, the existence of nonglycylable and nonglutamylable cortical tubulin isotypes. However, the simultaneous absence of the latter PTMs and of acetylation in
cortical microtubules would imply that at least some
a-tubulin isotypes possess sequence divergence both in
the carboxy-terminal and acetylation regions, as precisely
found in a digenean platyhelminth [Duvaux-Miret et al.,
1991] and in some other organisms [LeDizet and Piperno,
1987]. This assumption will be warranted only if a
diversity of tubulin isotypes is demonstrated in E. caproni,
both at the genetic and at the subcellular level.
An alternative situation would involve a specific
localization of the enzymes. Previous data about the
immunoreactivity of AXO 49 mAb and of a related
polyclonal Ab (PAT) in different systems such as Paramecium, quail oviduct and Drosophila testis [Adoutte et al.,
1991; Bressac et al., 1995; Fleury et al., 1995] have led to
postulation of an association of the glycylating enzyme to
the membrane. This hypothesis is not in agreement with
the present data, since in E. caproni the cortical microtubules that are lying in close contact with the membrane
are decorated with none of the antitubulin PTM antibodies tested, in contrast to the axonemal ones. A possibility,
then, is that the PTM enzymes might be specifically
associated with the axoneme itself.
Polyglycylation Discriminates Between Two
Axonemal Systems in the Same Organism
We have shown that the ciliary axonemes of E.
caproni protonephridia, possessing the classical 9 1 2
structure, are reactive with both TAP 952 and AXO 49
mAbs, indicating the presence of low and high levels of
Fig. 7. Double labelling immunofluorescence of spermatid microtubules with C105 (b-tubulin)
(a, d, g, j, m, p, s) and 6-11B-1 (acetylated tubulin) (b) or GT335 (glutamylated tubulin) (e, h, k, n) or
TAP 952 (glycylated tubulin) (q, t), and DNA staining (c, f, i, l, o, r), showing the sequential appearance of
PTMs during spermiogenesis in E. caproni. Bar 5 25 µm.
Tubulin Polyglycylation in Stable Microtubules
327
shows that evolutionary considerations may not be
the sole factor involved in the specification of the
glycylation level, contrary to suggestions from previous
studies.
In order to achieve the covalent binding of the
glycine residues, at least two distinct enzymes are expected to be required: one for the linkage of the first
glycine to a glutamate residue of the tubulin polypeptide
chain by a gCOOH-aNH2 amide linkage, and a second
for the formation of the aCOOH-aNH2 peptide bonds
between the added glycine residues of the lateral chain. In
this context, several types of mechanisms can be considered to account for these results.
The most simple explanation involves a differential
expression of the glycylating enzymes in the two cell
types of E. caproni: the first catalyzing the linkage of the
first Gly residue(s) to the tubulin polypeptide chain would
be simultaneously present in protonephridia and spermatozoa, whereas the second would be expressed in protonephridia only.
Another possibility concerns the regulation of the
enzymes. In this context, a reverse enzyme and/or
specific inhibitors, such as those previously reported for
tubulin acetylation [Maruta et al., 1986] and glutamylation [Audebert et al., 1993], could intervene in the
adjustment of the enzymatic equilibrium responsible for
the extent of glycylation in each compartment.
Therefore, the analysis of polyglycylation, by means
of two complementary antibodies, permits postulation of,
at least in E. caproni, a differential expression or regulation of the glycylating enzymes involved in two cell types
specialized in different functions.
In conclusion, whatever the mechanism involved
for segregating the right isoforms at the right place,
polyglycylation appears to act by two ways, either by its
absence/presence or by modulating the number of glycyl
units added to the tubulin molecule.
Fig. 8. Double labelling immunofluorescence of microtubules from
young and late spermatids of E. caproni, visible in the same field, with
C105 (b-tubulin) (a) and GT335 (glutamylated tubulin) (b); DNA
staining (c). Early (left) and late (right) spermatids show similar
intensities of labelling with C105 (a), but with GT335 (b) the labelling
intensity is stronger in late spermatids. A few mature strongly labelled
spermatozoa, randomly located in the field, can be observed. Bar 5
50 µm.
glycylation in this species. However, the lack of reactivity
of AXO 49 with the 9 1 ‘‘1’’ flagellar axonemes of
spermatozoa suggests the occurrence of only low levels
of glycylation in the latter cell. This last result is
reminiscent of that found in human spermatozoa [Bré et
al., 1996], contrasting with all other species tested, and
Sequential Occurrence of PTMs During Flagellar
Elongation Could Be Involved in the
Morphogenetic Clock of Spermiogenesis
The accessibility of the axonemal structure in E.
caproni flagella has permitted us to trace the appearance
of PTMs during spermiogenesis, both by means of
immunofluorescence and immunogold techniques.
It is noteworthy that the combined use of TAP 952
and AXO 49 is necessary to follow the time occurrence of
glycylation, instead of the exclusive use of AXO 49
[Bressac et al., 1995], specific to a restricted population
of glycylated isoforms. Indeed, in Paramecium, a lag
time is observed in the detection of the AXO 49 epitope,
in contrast to the TAP 952 one, in new microtubular
structures assembled in the course of cell division [Fleury
et al., 1995].
328
Iomini et al.
Fig. 9. Immunogold labelling of ultrathin sections of late spermatids of E. caproni with GT335
(glutamylated tubulin) (a–c) and TAP 952 (glycylated tubulin) (d–f). Randomly obtained transverse
sections at various levels of elongating spermatids were characterized according to Iomini and Justine
[1997]. For instance, the sections (c) and (f) are recognized as distal by the simultaneous presence of the
nucleus and a single axoneme. Bar 5 250 nm.
This study in Platyhelminthes shows that various
tubulin PTMs take place successively during the process
of flagellar elongation, that acetylation and thereafter
glutamylation appear precociously, and that these are
followed after a great delay by glycylation. The fact that
similar conclusions are reached from organisms belonging to evolutionarily very distant phyla—Platyhelminthes
(this report) and Arthropoda [Bré et al., 1996]—lead the
conclusion that this is a general phenomenon. The
sequential aspect of the PTM appearance is clearly
demonstrated in E. caproni. In this context, one might ask
whether acetylation and glutamylation are prerequisite
events for glycylation occurrence.
The late detection of the AXO 49 epitope in the
course of Drosophila spermiogenesis, at the time of the
individualization process, i.e., when the axonemal structure comes in contact with the membrane [Bressac et al.,
1995], has largely contributed to the hypothesis, discussed above, of the membrane localization of the
enzyme. Echinostoma caproni was expected to be a good
model to test this hypothesis because in this organism, the
individualization process does not exist. It is therefore
possible to dissociate the two events, namely, the coupling of the membrane to the axoneme and the time
occurrence of the glycylated epitopes. Indeed, from a
structural point of view, it exhibits an opposite situation
to that found in Drosophila during spermiogenesis: in the
young spermatid, comprising two free flagella (see Fig.
6), the membrane is close to the axoneme, whereas in the
late spermatid, after fusion of the three processes (Fig. 6),
only a few of the doublets are facing the membrane and
are not tightly placed against it [Iomini and Justine,
1997]. Thus the glycylated epitope appears in the late
stages even though the axoneme/membrane connection is
loose, whereas it is tight in the early stages.
In conclusion, the whole set of observations strengthens the previous hypothesis of a developmental regulation of glycylation occurrence during spermiogenesis
[Bré et al., 1996]. Moreover, it suggests that this event
might be involved in a morphogenetic clock for the end of
spermiogenesis rather than being simply the consequence
of a coupling of the membrane to the axoneme.
In addition to a possible morphogenetic role, glycylation has been postulated to be involved in sea urchin
spermatozoa motility [Bré et al., 1996]. When observed
out of the seminal vesicle, E. caproni spermatozoa are
able to beat but not to swim. As has been undertaken for
sea urchin spermatozoa, it would be worth investigating
whether TAP 952 is able to inhibit the flagellar beating.
Compared to sea urchin spermatozoa, which are deco-
Tubulin Polyglycylation in Stable Microtubules
rated with both AXO 49 and TAP 952 [Bré et al., 1996],
E. caproni spermatozoa are labelled with TAP 952
exclusively. This suggests that, in these sperm, axonemal
tubulin would posses an overall lower level of glycylation, possibly arranged in short chains. It may be asked
whether the structure of the glycyl chains could explain
the ability for the flagella to beat, but the inability for the
spermatozoa to swim. This would imply that the length of
the glycyl chain could influence, directly or by the way of
associated molecules, the swimming parameters of the
spermatozoon.
It is worth noting that the proximal appearance of
glycylated tubulin isoforms in the spermatids of E.
caproni is reminiscent of the proximo-distal decreasing
gradient of immunoreactivity observed in sea urchin
spermatozoa with AXO 49. Therefore, given the polar
growth of the axoneme, the extent of glycylation could be
considered, at least for some species, as a marker of the
oldest part of the flagellar axoneme. One might ask
whether these gradients merely reflect the age of the
consecutive parts of the axoneme, or provide means to
produce polarized associations of microtubules with
other molecules.
ACKNOWLEDGMENTS
The specimens were kindly provided by Sandrine
Trouvé and Annie Fournier (University of Perpignan,
France). C. I. acknowledges the help of Marina Allary for
editing early versions of the manuscript. Prof. B. G. M.
Jamieson kindly edited the English. This work was partly
supported by a BQR ‘‘Immunocytochimie des spermatozoı̈des de Plathelminthes et Nématodes’’ from the Muséum, the INTAS grant n° 93-2176, ‘‘Ultrastructure and
immunocytochemistry of the cytoskeleton of spermatozoa, eggs and fertilization in selected invertebrates species, for the understanding of phylogeny,’’ the CNRS, the
Université Paris-Sud, and the Muséum National d’Histoire
Naturelle. We are grateful to Prof. A. Adoutte for his
interest in this work. We thank the colleagues who
provided us with antibodies, Roselyne Tcheprakoff for
making the drawing, and L. Elu for help in photographic
work.
REFERENCES
Adoutte, A., Delgado, P., Fleury, A., Levilliers, N., Lainé, M.-C.,
Marty, M.-C., Boisvieux-Ulrich, E., and Sandoz, D. (1991):
Microtubule diversity in ciliated cells: evidence for its generation by post-translational modification in the axonemes of
Paramecium and quail oviduct cells. Biol. Cell 71:227–245.
Alexander, J.E., Hunt, D.F., Lee, M.K., Shabanowitz, J., Michel, H.,
Berlin, S.C., MacDonald, T.L., Sundberg, R.J., Rebhun, L.I.,
and Frankfurter, A. (1991): Characterization of posttranslational
modifications in neuron-specific class III beta-tubulin by mass
spectrometry. Proc. Natl. Acad. Sci. USA 88:4685–4689.
329
Arevalo, M.A., Nieto, J.M., Andreu, D., and Andreu, J.M. (1990):
Tubulin assembly probed with antibodies to synthetic peptides.
J. Mol. Biol. 214:105–120.
Audebert, S., Desbruyères, E., Gruszczynski, C., Koulakoff, A., Gros,
F., Denoulet, P., and Eddé, B. (1993): Reversible polyglutamylation of alpha- and beta-tubulin and microtubule dynamics in
mouse brain neurons. Mol. Biol. Cell. 4:615–626.
Barra, H.S., Arce, C.A., Rodriguez, J.A., and Caputto, R. (1974): Some
common properties of the protein that incorporate tyrosine as a
single unit and the microtubule proteins. Biochem. Biophys.
Res. Commun. 60:1384–1390.
Blose, S.H., Meltzer, D.I., and Feramisco, J.R. (1984): 10 nm filaments
are induced to collapse in living cells microinjected with
monoclonal and polyclonal antibodies against tubulin. J. Cell
Biol., 98:847–858.
Boucher, D., Larcher, J.C., Gros, F., and Denoulet, P. (1994): Polyglutamylation of tubulin as a progressive regulator of in vitro
interactions between the microtubule-associated protein Tau
and tubulin. Biochemistry 33:12471–12477.
Bré, M.-H., de Néchaud, B., Wolff, A., and Fleury, A. (1994):
Glutamylated tubulin probed in ciliates with the monoclonal
antibody GT335. Cell Motil. Cytoskeleton 27:337–349.
Bré, M.-H., Redeker, V., Quibell, M., Darmanaden-Delorme, J.,
Bressac, C., Cosson, J., Huitorel, P., Schmitter, J.-M., Rossier,
J., Johnson, T., Adoutte, A., and Levilliers, N. (1996): Axonemal
tubulin polyglycylation probed with two monoclonal antibodies: widespread evolutionary distribution, appearance during
spermatozoan maturation and possible function in motility. J.
Cell Sci. 109:727–738.
Bressac, C., Bré, M.-H., Darmanaden-Delorme, J., Laurent, M.,
Levilliers, N., and Fleury, A. (1995): A massive new posttranslational modification occurs on axonemal tubulin at the final step
of spermatogenesis in Drosophila. Eur. J. Cell Biol. 67:346–
355.
Callen, A.-M., Adoutte, A., Andreu, J.M., Baroin-Tourancheau, A., Bré,
M.-H., Ruiz, P.C., Clérot, J.-C., Delgado, P., Fleury, A.,
Jeanmaire-Wolf, R., Viklicky, V., Villalobo, E., and Levilliers,
N. (1994): Isolation and characterization of libraries of monoclonal antibodies directed against various forms of tubulin in
Paramecium. Biol. Cell 81:95–119.
Caron, J.M. (1997): Posttranslational modification of tubulin by
palmitoylation: I. In vivo and cell-free studies. Mol. Biol. Cell
8:621–636.
Daddow, L.Y.M. (1986): An abbreviated method of the double lead
stain technique. J. Submicrosc. Cytol. 18:221–224.
Duvaux-Miret, O., Baratte, B., Dissous, C., and Capron, A. (1991):
Molecular cloning and sequencing of the alpha-tubulin gene
from Schistosoma mansoni. Mol. Biochem. Parasitol. 49:337–
340.
Eddé, B., Rossier, J., Le Caer, J.-P., Desbruyères, E., Gros, F., and
Denoulet, P. (1990): Posttranslational glutamylation of alphatubulin. Science 247:83–85.
Eipper, B.A. (1974): Properties of rat brain tubulin. J. Biol. Chem.
249:1407–1416.
Fleury, A., Callen, A.-M., Bré, M.-H., Iftode, F., Jeanmaire-Wolf, R.,
Levilliers, N., and Clérot, J.-C. (1995): Where and when is
microtubule diversity generated in Paramecium? Immunological properties of microtubular networks in the interphase and
dividing cells. Protoplasma 189:37–60.
Fouquet, J.-P., Eddé, B., Kann, M.-L., Wolff, A., Desbruyéres, E., and
Denoulet, P. (1994): Differential distribution of glutamylated
tubulin during spermatogenesis in mammalian testis. Cell Motil.
Cytoskeleton 27:49–58.
Fried, B., and Huffman, J.E. (1996): The biology of the intestinal
trematode Echinostoma caproni. Adv. Parasitol. 38:311–368.
330
Iomini et al.
Fulton, C., and Simpson, P.A. (1976): Selective synthesis and utilisation of flagellar tubulin. The multitubulin hypothesis. In R.
Goldman, T. Pollard, and J. Rosenbaum (eds): ‘‘Cell Motility.’’
Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press,
pp. 987–1005.
Gagnon, C., White, D., Cosson, J., Huitorel, P., Eddé, B., Desbruyéres,
E., Paturle-Lafanechère, L., Multigner, L., Job, D., and Cibert,
C. (1996): The polyglutamylated chain of alpha-tubulin plays a
key role in flagellar motility. J. Cell Sci. 109:1545–1553.
Gelfand, V.I., and Bershadsky, A.D. (1991): Microtubule dynamics:
mechanim, regulation and function. Annu. Rev. Cell Biol.
7:93–116.
Greer, K., and Rosenbaum, J.L. (1989): Post-translational modifications of tubulin. In F.D. Warner and J.R. McIntosh (eds): ‘‘Cell
Movement, Vol. 2: Kinesin, Dynein and Microtubule Dynamics.’’ New York: Alan R. Liss, pp. 47–66.
Iomini, C., and Justine, J.-L. (1997): Spermiogenesis and spermatozoon of Echinostoma caproni (Platyhelminthes, Digenea): Transmission and scanning electron microscopy, and tubulin immunocytochemistry. Tissue & Cell 29:107–118.
Iomini, C., Raikova, O., Noury-Sraı̈ri, N., and Justine, J.-L. (1995):
Immunocytochemistry of tubulin in spermatozoa of Platyhelminthes. In B.G.M. Jamieson, J. Ausio, and J.-L. Justine (eds):
‘‘Advances in Spermatozoal Phylogeny and Taxonomy.’’ Mém.
Mus. Natn. Hist. Nat., 166:97–104.
Justine, J.-L. (1991): Phylogeny of parasitic Platyhelminthes: A critical
study of synapomorphies proposed on the basis of the ultrastructure of spermiogenesis and spermatozoa. Can. J. Zool. 69:1421–
1440.
Khan, I.A., and Ludueña, R.F. (1996): Phosphorylation of beta
III-tubulin. Biochemistry 35:3704–3711.
Kyhse-Andersen, J. (1984): Electroblotting of multiple gels: a simple
apparatus without buffer tank for rapid transfer of proteins from
polyacrylamide to nitrocellulose. J. Biochem. Biophys. Methods 10:203–209.
L’Hernault, S.W., and Rosenbaum, J.L. (1985): Chlamydomonas
alpha-tubulin is posttranslationally modified by acetylation on
the Sigma-amino group of a lysine. Biochemistry 24:473–478.
Laemmli, U.K. (1970): Cleavage of structural proteins during the
assembly of the head of bacteriophage T4. Nature 227:680–685.
LeDizet, M., and Piperno, G. (1987): Identification of an acetylation
site of Chlamydomonas alpha-tubulin. Proc. Natl. Acad. Sci.
USA 84:5720–5724.
LeDizet, M., and Piperno, G. (1991): Detection of acetylated alphatubulin by specific antibodies. Meth. Enzymol. 196:264–274.
Levilliers, N., Fleury, A., and Hill, A.-M. (1995): Monoclonal and
polyclonal antibodies detect a new type of post-translational
modification of axonemal tubulin. J. Cell Sci. 108:3013–3028.
Maruta, H., Greer, K., and Rosenbaum, J.L. (1986): The acetylation of
alpha-tubulin and its relationship to the assembly and disassembly of microtubules. J. Cell Biol. 103:571–579.
Mary, J., Redeker, V., Le Caer, J.-P., Promé, J.-C., and Rossier, J.
(1994): Class I and IVa beta-tubulin isotypes expressed in adult
mouse brain are glutamylated. FEBS Lett. 353:89–94.
Mary, J., Redeker, V., Le Caer, J.-P., Rossier, J., and Schmitter, J.-M.
(1996): Posttranslational modifications in the C-terminal tail of
axonemal tubulin from sea urchin sperm. J. Biol. Chem.
271:9928–9933.
Multigner, L., Pignot-Paintrand, I., Saoudi, Y., Job, D., Plessmann, U.,
Rüdiger, M., and Weber, K. (1996): The A and B tubules of the
outer doublets of sea urchin sperm axonemes are composed of
different tubulin variants. Biochemistry 35:10862–10871.
Ozols, J., and Caron, J.M. (1997): Posttranslational modification of
tubulin by palmitoylation: II. Identification of sites of palmitoylation. Mol Biol. Cell 8:637–645.
Paturle-Lafanechère, L., Eddé, B., Denoulet, P., Van Dorsselaer, A.,
Mazarguil, H., Le Caer, J.P., Wehland, J., and Job, D. (1991):
Characterization of a major brain tubulin variant wich cannot be
tyrosinated. Biochemistry 30:10523–10528.
Piperno, G., and Fuller, M.T. (1985): Monoclonal antibodies specific
for an acetylated form of alpha-tubulin recognize the antigen in
cilia and flagella from a variety of organisms. J. Cell Biol.
101:2085–2094.
Raff, E.C. (1994): The role of multiple tubulin isoforms in cellular
microtubule function. In J.S. Hyams and C.W. Lloyd (eds):
‘‘Microtubules.’’ New York: Wiley-Liss, pp. 85–109.
Redeker, V., Levilliers, N., Schmitter, J.-M., Le Caer, J.-P., Rossier, J.,
Adoutte, A. and Bré, M.-H. (1994): Polyglycylation of tubulin:
A posttranslational modification in axonemal microtubules.
Science 266:1688–1691.
Redeker, V., Melki, R., Promé, D., Le Caer, J.-P., and Rossier, J. (1992):
Structure of tubulin C-terminal domain obtained by subtilisin
treatment. The major alpha and beta tubulin isotypes from pig
brain are glutamylated. FEBS Lett. 313:185–192.
Richard, J. (1964): Trématodes d’oiseaux de Madagascar (Note III)
Espèces de la famille Echinostomatidae Poche 1926. Ann.
Parasitol. Hum. Comp. 34:607–620.
Rohde, K. (1990): Phylogeny of Platyhelminthes, with special reference to parasitic groups. Int. J. Parasitol. 20:979–1007.
Rüdiger, M., Plessman, U., Kloppel, K.D., Wehland, J., and Weber, K.
(1992): Class II tubulin, the major brain beta tubulin isotype is
polyglutamylated on glutamic acid residue 435. FEBS Lett.
308:101–105.
Rüdiger, M., Plessmann, U., Rüdiger, A.H., and Weber, K. (1995): Beta
tubulin of bull sperm is polyglycylated. FEBS Lett. 364:147–
151.
Schneider, A., Plessmann, U., and Weber, K. (1997): Subpellicular and
flagellar microtubules of Trypanosoma brucei are extensively
glutamylated. J. Cell Sci. 110:431–437.
Thompson, W.C. (1982): The cyclic tyrosination/detyrosination of
alpha tubulin. Meth. Cell Biol. 24:235–255.
Weber, K., Schneider, A., Müller, N., and Plessmann, U. (1996):
Polyglycylation of tubulin in the diplomonad Giardia lamblia,
one of the oldest eukaryotes. FEBS Lett. 393:27–30.
Wolff, A., de Néchaud, B., Chillet, D., Mazarguil, H., Desbruyères, E.,
Audebert, S., Eddé, B., Gros, F., and Denoulet, P. (1992):
Distribution of glutamylated alpha and beta tubulin in mouse
tissues using a specific monoclonal antibody, GT335. Eur. J.
Cell Biol. 59:425–432.
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