Annals of Botany 97: 679–693, 2006
doi:10.1093/aob/mcl023, available online at www.aob.oxfordjournals.org
INVITED REVIEW
Cytoskeleton and Morphogenesis in Brown Algae
C H R I S T O S K A T S A R O S *, D E M O S T H E N E S K A R Y O P H Y L L I S and B A S I L G A L A T I S
University of Athens, Faculty of Biology, Department of Botany, Athens 157 84, Greece
Received: 25 September 2005 Returned for revision: 5 November 2005 Accepted: 28 November 2005 Published electronically: 8 February 2006
Background Morphogenesis on a cellular level includes processes in which cytoskeleton and cell wall expansion
are strongly involved. In brown algal zygotes, microtubules (MTs) and actin filaments (AFs) participate in polarity
axis fixation, cell division and tip growth. Brown algal vegetative cells lack a cortical MT cytoskeleton, and are
characterized by centriole-bearing centrosomes, which function as microtubule organizing centres.
Scope Extensive electron microscope and immunofluorescence studies of MT organization in different types of
brown algal cells have shown that MTs constitute a major cytoskeletal component, indispensable for cell
morphogenesis. Apart from participating in mitosis and cytokinesis, they are also involved in the expression
and maintenance of polarity of particular cell types. Disruption of MTs after Nocodazole treatment inhibits cell
growth, causing bulging and/or bending of apical cells, thickening of the tip cell wall, and affecting the nuclear
positioning. Staining of F-actin using Rhodamine-Phalloidin, revealed a rich network consisting of perinuclear,
endoplasmic and cortical AFs. AFs participate in mitosis by the organization of an F-actin spindle and in cytokinesis
by an F-actin disc. They are also involved in the maintenance of polarity of apical cells, as well as in lateral branch
initiation. The cortical system of AFs was found related to the orientation of cellulose microfibrils (MFs), and
therefore to cell wall morphogenesis. This is expressed by the coincidence in the orientation between cortical AFs
and the depositing MFs. Treatment with cytochalasin B inhibits mitosis and cytokinesis, as well as tip growth of
apical cells, and causes abnormal deposition of MFs.
Conclusions Both the cytoskeletal elements studied so far, i.e. MTs and AFs are implicated in brown algal cell
morphogenesis, expressed in their relationship with cell wall morphogenesis, polarization, spindle organization and
cytokinetic mechanism. The novelty is the role of AFs and their possible co-operation with MTs.
Key words: Actin, cytoskeleton, microtubules, morphogenesis, Phaeophyceae, polarity, vegetative cells.
INTROD UCTION
Morphogenesis, according to the etymology of the term
[Greek morphi (= shape) + genesis (= creation)], is a highly
controlled process by which the enormous variability of
shapes of organisms arise. Since the first research studies
in the plant kingdom, the great variety of forms occurring
in algae made them a precious source of model systems for
basic research of both morphological and physiological
importance.
Brown algae constitute a large group, which occupies a
particular position in the evolutionary process. Their unique
characteristics distinguish them from other classes within
Heterokontophyta. The brown algal cell also shows particular characteristics in structure, cytoskeleton organization
and division process. All the above make brown algae a
very interesting model for many morphogenetical studies
(Katsaros, 1995, 2001).
The role of the cytoskeleton in cell wall morphogenesis is
well established in higher plants (Fowler and Quatrano,
1997; Baskin, 2001; Hasezawa and Kumagai, 2002;
Smith, 2003; Wasteneys, 2004). Cortical microtubules
(MTs) have been shown to determine directly (Burk and
Ye, 2002) or indirectly microfibril (MF) orientation in
higher plant cells, possibly defining the distribution of cellulose synthesizing enzymes in the plasmalemma (Tsekos,
1999; Baskin, 2001; Hasezawa and Kumagai, 2002). This is
also the case in algae possessing cortical MTs (Mizuta,
1992; Tsekos, 1999). Contrary to the above, brown algal
* For correspondence. E-mail [email protected]
cells lack a cortical MT cytoskeleton and are characterized
by centriole-bearing centrosomes, which function as the
microtubule-organizing centres for all cell MTs (Katsaros
et al., 1983; Katsaros and Galatis, 1992; Motomura, 1994).
Extensive electron microscope and immunofluorescence
studies of MTs in different types of brown algal cells
have shown that they play a primary role in their function.
Apart from participating in mitosis, they also play a key role
in cytokinesis, as well as in the expression of polarity of
particular cell types (Katsaros et al., 1983; Katsaros, 1992;
Katsaros and Galatis, 1992; Kropf, 1992a, b, 1994;
Karyophyllis et al., 1997). The interactions between plasma
membrane with cytoskeleton and cell wall have been
reviewed by Fowler and Quatrano (1997; see also
Baluska et al., 2000, 2001). In addition to MTs, the role
of F-actin cytoskeleton in morphogenetic processes of
brown algae, such as polarization, cell cycle, and cell
wall expansion has recently been studied in more detail
(Karyophyllis et al., 2000a, b; Katsaros et al., 2002;
Hable et al., 2003; Bisgrove and Kropf, 2004; Hable and
Kropf, 2005).
The aim of the present review is to summarize the
available data on the role of the cytoskeleton in morphogenesis of brown algae. The organization and function of
the cytoskeleton in fucoid zygotes has been repeatedly
reviewed (Kropf et al., 1998, 1999; Hable et al., 2003),
therefore, in this paper, special attention is given to the
role of MTs and actin filaments (AFs) in morphogenesis
of vegetative cells. This topic, as far as is known, has not
been reviewed so far.
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Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
680
A
B
C
D
F I G . 1 Apical cells of Sphacelaria. (A) Light microscope image of two apical cells of S. rigidula. (B) TEM image of a dividing apical cell of S. tribuloides.
(C) Immunofluorescence labelling of tubulin in apical cell of S. rigidula. (D) Cortical F-actin organization in apical cell of S. rigidula (from Karyophyllis et al.,
2000b). Scale bars: A = 20 mm; B = 10 mm; C = 5 mm; D = 5 mm.
POLAR ORGANIZATION OF THE
P R O T O P L A S T : TI P G R O W T H
Microtubule cytoskeleton
Cells frequently exhibit asymmetric distribution of organelles, proteins or cytoskeletal components along a particular
axis. This internal organization is referred to as ‘cell
polarity’ (Cove, 2000). Polarized cells exhibit different
morphological and/or molecular characteristics at opposite
ends aligned along an axis (Grebe et al., 2001; Baluska et al.,
2003). The use of fucalean zygotes as models for the study
of polarization mechanisms started very early (Knapp,
1931; Jaffe, 1958; Nuccitelli, 1978) and still continues
(Kropf, 1994, 1997; Kropf et al., 1998; Brownlee and
Bouget, 1998; Hable and Kropf, 1998, 2000, 2005;
Robinson et al., 1999; Belanger and Quatrano, 2000a;
Brownlee et al., 2001; Hable et al., 2003; Bisgrove and
Kropf, 2004). The ultrastructural examination of the unfertilized egg revealed a symmetrical distribution of cell
elements (Brawley et al., 1976a, b). The first signs of polarization, which is induced by environmental cues, become
visible 8–10 h after fertilization by the secretion of adhesive
material at the future growth site (Quatrano, 1990; Kropf,
1992a, b, 1997; Goodner and Quatrano, 1993; Hable and
Kropf, 1998, 2000). In the apolar zygote the MT cytoskeleton consists of numerous MTs initially non-centrosomal,
directing from the perinuclear cytoplasm, and later from the
centrosome towards the cell periphery, without any particular orientation. By the time of rhizoid emergence, MTs are
focused on two centrosomes, the axis of which is perpendicular to the direction of the polarity axis. Intense MT
bundles extend from each centrosome to the rhizoid pole.
After the emergence of the outgrowth at the rhizoid site, the
centrosome–nucleus system rotates by 90 , thus becoming
axially oriented (Kropf et al., 1990; Allen and Kropf, 1992;
Kropf, 1992a, b, 1994). Depolymerization of MTs does not
affect zygote polarization or rhizoid emergence, meaning
that MTs are not directly implicated in the establishment of
polarity in fucalean zygotes (Brawley and Quatrano, 1979;
Kropf et al., 1989). However, the asymmetrical division that
follows is critical for the future development of the young
embryo. Experimental alteration of the plane of this division
leads to the formation of abnormal embryos and affects their
viability (Bisgrove and Kropf, 1998). It is interesting that in
a recent paper, Corellou et al. (2005), using microinjection
of fluorescent tubulin into living Fucus zygotes, found a
distinct cortical MT array that is reorganized during polarization and rhizoid emergence. According to these authors
this Mt configuration is crucial for the regulation of rhizoid
growth. However, a similar MT organization was never
reported in any of the earlier studies dealing with MT organization in fucalean zygotes (Brawley et al., 1976a, b; Kropf
et al., 1990; Allen and Kropf, 1992; Bisgrove and Kropf,
1998, 2001, 2004; Nagasato et al., 1999b).
Contrary to the zygotes of Fucales, the tubular apical cells
of Sphacelariales display a permanent polarity, clearly visible at a light microscope level (Fig. 1A). This polar organization is directly related to the particular growth pattern of
these cells (Katsaros, 1980, 1995; Katsaros et al., 1983),
which is similar to that of tip-growing cells like fungal
hyphae, root hairs, pollen tubes, etc. It represents a gradual
distribution of the cell elements along the longitudinal axis
of the cell (Fig. 1B). The tip region is occupied by a large
number of endoplasmic reticulum (ER) membranes associated with active dictyosomes and relatively small vacuoles,
while the basal area is highly vacuolated. In this organelle
gradient the nucleus occupies a particular position. The
apical cell wall is continuously extending, and consists of
two layers, a thin external one consisting of amorphous
material, and an internal with a few randomly oriented
681
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
a
L1
L2
b
L3
c
L4
d
e
A
B
C
F I G . 2. Drawings showing the MT, AF and MF organization in the apical cell of S. rigidula. (A) MTs are radiating out of the centrosomes. More thin MT
bundles are directed towards the apex, while fewer thick ones are directed towards the base of the cell. (B) Pattern of cortical AF arrangement along the axis of
the cell. (C) Cell wall layers (L1, L2, L3 and L4) and MF orientation in different areas (a–e) along the apical cell (from Karyophyllis et al. 2000b). The
rectanglar area towards the top of the left side of the cell is shown magnified.
MFs embedded in an amorphous matrix (Karyophyllis et al.,
2000b). Subunits of cellulose synthases or entire terminal
complexes (TCs) are probably transported via dictyosome
vesicles to the plasmalemma of the tip region (Katsaros
et al., 1996; Reiss et al., 1996). Recent freeze-fracture studies of apical cells of Syringoderma phinneyi have revealed
an apico-basal gradient of TC distribution, with a higher
density of TCs in the apical part, where more intense
cellulose synthesis takes place. This seems to reflect the
tip growth of the apical cells, suggesting a relationship
between cellulose synthesis with the growth pattern
(Schüssler et al., 2003).
Immunofluorescence labelling of tubulin showed that the
interphase MT system in apical cells of Sphacelaria rigidula is strongly polarized. It extends from the two centrosomes towards the cortical cytoplasm, and forms a fine
meshwork. The apical dome of the cell is traversed by
numerous thin MT bundles, while the basal part is traversed
by a few thick ones meandering among the vacuoles
(Katsaros, 1992; Figs 1C and 2A). It can be assumed that
they stabilize and mechanically support the cytoplasmic
strands that are probably formed by the acto-myosin complex (see also Panteris et al., 2004). The plus ends of MTs in
the apical cytoplasm frequently reach and probably contact
the plasma membrane, where sometimes they almost curve
(Fig. 1C). In animal and yeast cells they seem to interact
with the cell cortex via plus end MT-associated proteins,
like +TIPs (e.g. CLIPs, CLASPs, cytoplasmic dynein/
dynactin, EB1, etc.; Mimori-Kiyosue and Tsukita, 2003).
These proteins act mainly as microtubule-stabilizing factors
and at the same time often link MTs to various cellular
structures, such as the cell cortex (Carvalho et al., 2003;
Bisgrove et al., 2004; Gundersen et al., 2004; Akhmanova
and Hoogenraad, 2005). The MT tip proteins interact with
bridging proteins that guide the MTs to cortical receptors
driven by AFs (Gundersen et al., 2004). On the other hand,
interactions between MT tips and motors can cause translocation of vesicles along MTs. Therefore MT plus ends can
serve as cargo-loading sites for minus-end-directed transport (Vaughan et al., 2002), or deliver cargos to particular
cell regions (Felerbach et al., 2004). Similar activities can
be attributed to the MTs of brown algal cells that bear
centrosomes like animal cells, and to the MTs approaching
the tip of the apical cell of S. rigidula where expansion takes
plase. Examination of the MT organization during mitosis
and cytokinesis revealed that the above polar organization
of MTs in apical cells is obvious during their whole cell
cycle (Katsaros, 1992).
Treatment with taxol induced dramatic changes in MT
organization in both interphase and dividing apical cells of
S. rigidula. Massive assembly of MTs occurs in the cortical
and subcortical cytoplasm of interphase cells out of the
centrosome areas. Mitosis and cytokinesis were inhibited
and numerous MTs were found in the perinuclear cytoplasm
of mitotic cells. It has been suggested that centrosome
dynamics in MT nucleation varies during the cell cycle,
and that in case centrosome activity is disturbed, the cortical/subcortical cytoplasm assumes the ability to assemble
MTs (Dimitriadis et al., 2001).
Tubulin immunolocalization in isolated protoplasts of
apical cells of Sphacelaria sp. revealed a different pattern
of MT organization, both in interphase and during the first
division. The cell becomes apolar and the MTs diverge
symmetrically from the centrosomes towards the cell cortex, during wall regeneration (Rusig et al., 1993, 1994). The
loss of polarity in MT organization after removal of the cell
wall is retained for at least the first divisions of the regenerating protoplasts. This fact underlines the role of the cell
wall in polarization of brown algal cells (Quatrano and
Shaw, 1997; Ouichou and Ducreux, 2000; Bisgrove and
Kropf, 2001). The polarity is re-established with the
initiation of a new apical cell from a mass of about ten
682
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
undifferentiated cells (Ducreux and Kloareg, 1988). Similar
behaviour characterizes the cytoskeleton of young and
regenerating protoplasts of Macrocystis pyrifera gametophytes (V. Varvarigos, unpubl. res).
After the first division of the germinating fucalean
zygote, the rhizoid cell behaves like a tip-growing cell.
The nature of tip growth is first manifested in wall structure.
The previously uniform external cell wall becomes thinner
at the tip compared with subapical regions (Allen and Kropf,
1992; Kropf et al., 1992). Experimental depolymerization
of MTs using drugs like oryzalin, ami-prophos methyl, or
Nocodazole (Nz) does not affect the fixation of the polar
axis and the rhizoid germination in fucalean zygotes
(Quatrano, 1973; Brawley and Quatrano, 1979; Kropf
et al., 1990; Kropf, 1992a, b, 1994). However, MTs are
involved in rhizoid morphogenesis, as in other tip-growing
cells, e.g. moss protonemata (Doonan et al., 1985, 1988) or
fungal hyphae (Raudaskoski et al., 1994; Robertson and
Vargas, 1994; Rupes et al., 1995). When MTs are destroyed
by anti-MT agents, the advanced developmental events are
severely inhibited and the zygotes form aberrant rhizoids
(Kropf, 1994). Similarly, in S. rigidula apical cells,
treatment with Nz disturbs polarity and affects the growth
pattern. The distribution of the organelles and particularly
dictyosomes becomes more uniform, and the cell wall of the
tip region appears thicker, while the cell sometimes bends.
This suggests that MTs are probably involved in the transport of vesicles with wall materials to the tip region
(Karyophyllis et al., 1997; Karyophyllis, 2003). Nagasato
and Motomura (2002b) reported that Golgi-derived vesicles
are transported along MTs towards the division plane of
cytokinetic cells of Scytosiphon lomentaria. A detailed
overview on the MT cytoskeleton in tip-growing plant
cells is presented in a number of recent reviews (Mathur
and Hulskamp, 2002; Sieberer et al., 2005; Smith and
Oppenheimer, 2005).
MTs in tip-growing cells are also involved in the correct
positioning of the nucleus. In most tip growing cells, MT
depolymerization disturbs the nuclear positioning
thus affecting the growth pattern (Doonan et al., 1985;
Lloyd et al., 1987; Joos et al., 1994; Heath et al., 2000).
Similar results have been obtained in apical cells of
S. rigidula treated with Nz (Karyophyllis et al., 1997;
Karyophyllis, 2003).
Actin filament cytoskeleton
The role of actin in establishment and maintenance of cell
polarity has been well documented. The asymmetric spatial
distribution and activity of the actin cytoskeleton, and the
conserved signalling molecules that regulate it, play a role
in cell polarity in metazoans, fungi (Johnson, 1999), amoebozoans (Chung et al., 2000) and higher plants (Yang,
2002). GTPases in the Ras superfamily, especially ones
in the Rho family are such signalling molecules that regulate
the actin cytoskeleton (Etienne-Manneville and Hall, 2002).
Recently, cDNA that encodes a Rho family GTPase was
isolated from Fucus distichous (Fowler et al., 2004). The
small GTPase encoded (FdRac1) displays a polarized localization, being concentrated close to the growing tip of the
rhizoid. This localization together with the evidence for
some functional overlap with yeast Cdc42p, support
the hypothesis that FdRac1 regulates algal cell polarity,
possibly via actin cytoskeleton (Fowler et al., 2004).
In tip-growing cells of higher plants and fungi showing
permanent polarity, like pollen tubes, root hairs or fungal
hyphae, the actin cytoskeleton supports the polar architecture of the cytoplasm and determines the direction of cell
expansion (Heath, 1990, 2000; Steer, 1990; Pierson and
Cresti, 1992; Derksen et al., 1995; Miller et al., 1999;
Emons and de Ruijter, 2000; Geitman and Emons, 2000;
Vidali and Hepler, 2000; Hepler et al., 2001; Wasteneys and
Galway, 2003). AFs traverse the cytoplasm of these tubular
cells in an axial direction, possibly facilitating the transport
of vesicles to the tip. However, actin is not always present at
the very tip region of root hairs and pollen tubes (Miller
et al., 1996, 1999; Emons and de Ruijter, 2000; Vidali and
Hepler, 2000; Lovy-Wheeler et al., 2005), while in developing fungal hyphae a dense meshwork of AFs is usually
present at the tip (Heath, 1990, 2000). It must be noted that
disruption of actin inhibits tip-growth in both root hairs and
pollen tubes (Gibbon et al., 1999; Miller et al., 1999;
Baluska et al., 2000).
The involvement of F-actin in the establishment of polarity in zygotes of Fucus or Pelvetia has been repeatedly
underlined. Early studies using cytochalasin B (CB) have
indicated that F-actin is involved in the conversion of a light
gradient to a morphological gradient in these cells (Nelson
and Jaffe, 1973; Quatrano, 1973). More recent studies using
anti-actin agents like cytochalasin or lantrunculin further
support the hypothesis that photopolarization of fucalean
zygotes is actin-dependent (Hable and Kropf, 1998;
Robinson et al., 1999; see also review in Fowler and
Quatrano, 1997). During all these years enough efforts
were made to visualize AFs (Brawley and Robinson,
1985; Kropf et al., 1989), since these fine structures are
not well preserved after chemical fixation. These difficulties
are commonly attributed to the high sensitivity of F-actin to
aldehyde fixatives (Doris and Steer, 1996; Miller et al.,
1996). However, more recent findings showed that rather
than aldehyde fixation, some further steps in the procedures
used for actin visualization are critical for preserving
F-actin (Vitha et al., 2000). To overcome this problem,
new approaches were developed, by introducing fluorescent
phalloidin into living cells (Alessa and Kropf, 1999; Pu
et al., 2000). These techniques have shown that F-actin
is organized into cortical patches, which during photopolarization are localized at the rhizoid pole. In the young
germinating rhizoid, these patches become a ring-like
configuration that is located in the subapical zone of the
elongating tip (Alessa and Kropf, 1999; Pu et al., 2000).
Disruption of F-actin using lantrunculin B blocks photopolarization, probably by inhibiting the formation of cortical Ca2+ gradients (Pu et al., 2000). However, in all the
above studies the staining usually revealed fluorescent
patches, but not AFs.
Contrary to the zygotes of Fucus and Pelvetia, which are
apolar cells with uniform distribution of cell elements and
become polarized some time after fertilization, the
apical cells of some filamentous brown algae like the genera
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
Sphacelaria, Halopteris, Syringoderma, etc., show an inherent cellular polarity. This is closely related to the growth
pattern of these cells and is strongly supported by the
particular organization of the cytoskeletal elements. The
application of a modified protocol for F-actin staining
with Rhodamine-Phalloidin (Rh-Ph), in vegetative cells
of a variety of brown algal species, allowed the observation
of clear images of AFs, and thus the detailed examination of
their organization (Karyophyllis et al., 2000a). It was found
that vegetative cells of brown algae bear a well-organized
cytoskeleton of AFs, consisting of cortical (Figs 1D and
2B), perinuclear and endoplasmic arrays of AFs. The
perinuclear and the endoplasmic AFs are organized in a
similar way in almost all the cell types examined. However,
the cortical AFs show variable arrangements in different
types of cells. In the tip-growing tubular apical cells of
S. rigidula their organization is quite complicated. The
dome area of the cell is traversed by short and thin bundles
of AFs in a rather random distribution. At the base of the
hemispherical part, AFs form a ring that is perpendicular to
the cell longitudinal axis. Below this area the cell becomes
tubular and the cortical AFs show a longitudinal or helical
AF arrangement, parallel to the cell axis (Figs 1D and
2B) (Karyophyllis et al., 2000a). The above organization
is involved in the formation of the polar growth pattern
(tip growth), by controlling cell wall morphogenesis
(Karyophyllis et al., 2000b; see below). The nucleation
mechanism of all the above dynamic actin arrays is not
known. In this direction, the role of the highly conserved,
actin-nucleating, Arp2/3 complex (Deeks and Hussey,
2003; Dos Remedios et al., 2003) was investigated recently
in Silvetia compressa (Hable and Kropf, 2005). A subunit of
the complex (Arp2) was immunolocalized in zygotes, and
the data indicate that the Arp2/3 complex participates in
nucleating all the new actin arrays, but not the cytokinetic
actin disc (Hable and Kropf, 2005).
The participation of the F-actin cytoskeleton in the
maintenance of polarity of apical cells of S. rigidula was
confirmed by experiments in which F-actin was affected by
CB. After 24 h incubation in the drug, the growth of the
apical cell was inhibited and its polar organization was
severely disturbed. The organelles were evenly distributed
along the cell axis and the wall of the apical region became
thicker than in normal cells. This allows the hypothesis that
AFs are involved in the morphogenesis of tip-growing cells,
alone or in co-operation with MTs and associated proteins.
The interaction of dynamic, growing MTs with AF arrays
associated with the cell cortex, their mutual regulation and
their participation in morphogenetic processes, have been
described in a number of different animal cell types (Goode
et al., 2000; Gundersen, 2002).
BRAN CH FORMATION
Microtubule cytoskeleton
In some filamentous brown algae like S. rigidula or Ectocarpus siliculosus lateral branches are formed in differentiating or even differentiated thallus areas. The process of
branching starts with a local outgrowth of the external cell
683
wall which soon becomes thin in the tip region and forms a
dome-like protrusion similar to that of tubular apical cells
(Katsaros, 1980, 1995). During branch initiation MT
cytoskeleton undergoes significant alterations (Katsaros,
1992). At first the centrosome–MT system rotates and the
axis determined by the two centrosomes becomes oblique to
the thallus axis. At a second stage, the nucleus migrates
towards the wall outgrowth. A rich meshwork consisting
of thin MT bundles radiating from the proximal centrosome
extends towards the tip of the nascent branch. On the
contrary, thick MT bundles radiate out of the distal centrosome traversing the cytoplasm among vacuoles of the basal
part. This organization indicates the establishment of a new
polarity axis in a more or less differentiated thallus cell. The
MT cytoskeleton is involved in this polarization process by
changing the positioning of the nucleus and possibly participating in the local softening of the cell wall, probably by
transportation and exocytosis of some loosening factor via
dictyosome vesicles. The subsequent cell division separates
the apical cell of the young branch, which then behaves like
the main apical cell of the thallus (Katsaros, 1992, 1995).
Similar MT reorganization has been observed in cells of
Macrocystis pyrifera gametophytes which give lateral
branches. However, in this case the branch cell separated
by the first division does not behave as an apical cell
(Varvarigos et al., 2005).
Actin filament cytoskeleton
F-actin seems to be involved in the process of branch
formation. Before any sign of protoplast germination or
thallus branching of M. pyrifera gametophytes, radial AF
configurations are organized in the prospective sites of the
wall outgrowth. These initially pointed radial AF structures
change to radial-circular configurations located at the base
of the emerging protrusion, while at more advanced stages
they disappear. Treatment with latrunculin B resulted in the
inhibition of germination and branching, suggesting that
these radial AF configurations are involved in cell polarization (Varvarigos et al., 2004; Fig. 3). Similarly in
S. rigidula, AFs are gathered in the sites of wall protrusion
during branch formation. It is speculated that some motor
protein(s) travel(s) along these actin structures transporting
many types of cargo, including secretory vesicles. Polarized
secretion of vesicles carrying cell wall remodelling
enzymes and new cell wall constituents may promote
local cell wall deformation and branch emergence. As
described above, the first sign of branching is the formation
of a wall outgrowth in a rather differentiated cell of the
thallus. Rh-Ph staining revealed an increased F-actin fluorescence at the site of the bulge. At a later stage, when the
outgrowth became a short tube with a dome-like tip, this
F-actin reorganized and a dense fluorescence was observed
at the base of the dome (Karyophyllis, 2003). This F-actin
system is similar to that described by Allessa and Kropf
(1999) (see also Kropf et al., 1998) and Pu et al. (2000) in
germinating fucalean zygotes. It seems likely that AFs are
involved in the establishment of a new polarity axis during
branch formation as in apolar zygotes. The F-actin ring that
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Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
b1
c1
a
b2
c2
F I G . 3. Diagrammatic representation of the AF organization in
regenerating protoplasts of M. pyrifera female gametophytes. a, AF organization in young protoplasts and protoplast-derived cells. b, Pointed-radial
actin configurations prior to the formation of the protrusion. c, Radialcircular actin configurations and emergence of the protrusion. In b and c
two views are shown: 1, from the top; 2, from the side.
is formed at a later stage is involved in the morphogenesis of
the tip-growing cell, and resembles the ring found in apical
cells of S. rigidula. Similar F-actin rings have also been
described in fern protonemata (Kadota and Wada, 1989;
Quader and Schnepf, 1989) and fungal hyphae
(Bachewich and Heath, 1998). The function of these rings
is not yet completely understood. For those found in
germinating fucoid zygotes it has been hypothesized
that they are implicated in tip reinforcement in the boundary between the fragile apex and the subapical rhizoid
domain (Kropf et al., 1998; Alessa and Kropf, 1999). In
this region a ring of tight adhesion between cortical
F-actin, plasma membrane and the cell wall has been
observed in Pelvetia compressa rhizoids (Henry et al.,
1996). It has also been proposed that this actin ring
participates in the maintenance of Ca2+ ion pumps in the
rhizoid tip (Pu et al., 2000). A different role of a transverse
AF ring in cell wall morphogenesis of tip-growing cells
is reported in apical cells of S. rigidula, where AFs
have been found related to the orientation of the MFs
(Karyophyllis et al., 2000b; Karyophyllis, 2003). They
are probably related to the formation and maintenance of
the tubular form of the cell.
CELL CYCLE
Microtubule cytoskeleton
The absence of cortical MTs, MT preprophase band and
phragmoplast, as well as the presence of centrosomes in
brown algal cells make them a quite interesting model
for the study of MT organization during the cell cycle.
However, the difficulties in the preservation of the fine
structure in transmission electron microscope (TEM) studies resulted in an incomplete description of the organization
of MTs in interphase and dividing brown algal cells
(Neushul and Dahl, 1972; Galatis et al., 1973, 1977;
Katsaros et al., 1983; Katsaros and Galatis, 1985, 1988,
1990; see also Katsaros, 1980, 1995). The particular composition of their cell wall also caused a considerable delay in
the successful application of immunofluorescence staining
of tubulin and in the detailed analysis of MT organization
(Katsaros, 1992; Katsaros and Galatis, 1992; Rusig et al.,
1993, 1994; Karyophyllis et al., 1997; Katsaros and Sala,
1997).
The organization of MTs during mitosis shows some
differences between tip-growing apical cells, like those
of S. rigidula, and other types of cells with more diffuse
growth, like subapical cells of S. rigidula, or apical cells of
Dictyota dichotoma (Katsaros, 1992; Katsaros and Galatis,
1992; Rusig et al., 1993, 1994). In tip-growing cells this
becomes evident by the continuous coincidence between
cytoplasmic polarity axis and the axis of the MT–
centrosome system, as well as by the difference in the
dynamics of the MTs of the apical part compared with
those of the basal one. The latter is evident by the fact
that the apical MTs disappear last with the entrance of the
cell in mitosis and reappear first on anaphase (Katsaros,
1992, 1995; Rusig et al., 1994, 2001; Karyophyllis et al.,
1997).
By the completion of anaphase, the spindle reaches its
longest size, therefore the daughter nuclei are formed at a
distance from each other, which is longer in elongated cells.
In brown algal vegetative cells, contrary to higher plants,
cytokinesis starts long after the completion of telophase. In
the meantime daughter nuclei assume a completely
interphase appearance (Katsaros, 1980, 1992, 1995;
Katsaros et al., 1983; Katsaros and Galatis, 1992). At
this stage spindle MTs are depolymerized and cytoplasmic
MTs reassemble from the two oppositely placed centrosomes. The daughter nuclei approach each other and the
‘cytokinetic’ MTs form two cage-like configurations, surrounding the daughter nuclei and overlapping at the midplane (Katsaros and Galatis, 1992). At a later stage the
daughter nuclei move apart again, probably by interaction
of the two interdigitating MT systems via motor proteins. In
this way, two nuclear-cytoplasmic domains (NCDs) or
‘cytoplasts’ are organized (Porter and McNiven, 1982;
Porter et al., 1983; Pickett-Heaps et al., 1999). In transverse
divisions of elongated cells, a clear space free of MTs is
formed between the two NCDs. The cytokinetic diaphragm
is then developed in the mid-plane between the two NCDs,
by the fusion of vesicles and cisternae (Katsaros and Galatis,
1992; Nagasato and Motomura, 2002b). The NCD concept
was described, although not termed, in sporangia of
Halopteris filicina and in dividing vegetative cells of
D. dichotoma (Katsaros and Galatis, 1986, 1992). Treatment with anti-MT or anti-actin agents inhibits cytokinesis,
meaning that both MTs and AFs are involved in this process
(Karyophyllis et al., 1997, 2000a; Bisgrove et al., 2003).
However, although the study of cell division in brown algae
started quite early, the cytokinetic mechanism is still a
matter for discussion (Galatis et al., 1973; Rawlence,
1973; Markey and Wilce, 1975; Brawley et al., 1977;
LaClaire, 1981; Katsaros et al., 1983; Belanger and
Quatrano, 2000b; Nagasato and Motomura, 2002b;
Bisgrove and Kropf, 2004).
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
Actin filament cytoskeleton
The organization of AF cytoskeleton was studied in
vegetative cells of S. rigidula during the whole cell cycle
(Karyophyllis et al., 2000a; Karyophyllis, 2003). The
interphase organization of AFs was described above. During
mitosis the F-actin cytoskeleton is completely reorganized.
The cortical F-actin system remains well organized until
telophase. At prophase the fluorescence of the endoplasmic
AFs becomes weak, and the perinuclear AFs predominate.
In parallel, a bipolar spindle of AFs starts forming, that is
disorganized after anaphase. Although at these stages the
filamentous character of AFs is not always visible, it is clear
that their organization resembles that of spindle MTs. The
poles of the F-actin spindle coincide with those of the MT
spindle but they cover a more broad area. At metaphase,
AFs extend from the poles towards the chromosomes, and
during anaphase interzonal AFs are visible. By the progress
of telophase, the daughter nuclei move apart from each
other. The perinuclear AF system is still intense, but the
fluorescence in the interzonal area temporarily weakens. At
advanced telophase the interzonal AF system becomes
gradually more intense, and when the two daughter nuclei
have gained an interphase organization, all the other F-actin
systems are gradually reduced, except for the interzonal
one. Finally, before any sign of cytokinetic diaphragm
formation, a complete, dense F-actin disc is formed in the
plane of the future division. The assembly of an actin disc
midway between daughter nuclei was also confirmed in
fucoid zygotes (Bisgove and Kropf, 2004). It must be
noted here that, in vacuolated cells of M. pyrifera gametophytes, a unique pattern of F-actin organization participates
in the control of cytokinesis. This involves an F-actin ring
organized in the cortical cytoplasm which develops inwards
to form the F-actin disc (Varvarigos et al., 2005). TEM
examination of similar stages revealed vesicles and cisternae gathering in the area of the F-actin disc. Fusion of these
patches gives rise to the membranous diaphragm separating
the daughter cells. Although a centrifugal development of
the diaphragm has been proposed in zygotes of brown algae
(Nagasato and Motomura, 2002b; Bisgorve and Kropf,
2004), this seems not to be the case in vegetative cells
examined so far (see also Karyophyllis et al., 2005a).
The above data show that F-actin is involved in mitosis
and cytokinesis of brown algal vegetative cells, possibly cooperating with spindle and cytokinetic MTs. Evidence in
favour of this hypothesis was provided by experiments
using CB. Cells treated with CB at a premitotic stage did
not complete mitosis, but were blocked at a stage resembling metaphase. The absence of anaphase stages after treatment implied that AFs possibly participate in chromosome
movement. Similar results have been reported by Forer
(1985, 1988) in animal cells, where disruption of AFs by
cytochalasin also inhibited cytokinesis. The mechanism of
cytokinesis in brown algal cells is not yet fully understood.
However, a possible scenario would be drawn by correlation
of the above results with the available ultrastructural and
MT-immunofluorescence data (Galatis et al., 1973;
Rawlence, 1973, Markey and Wilce, 1975; Brawley et al.,
685
1977; LaClaire, 1981; Katsaros et al., 1983; Katsaros, 1992;
Katsaros and Galatis, 1992; Karyophyllis et al., 1997;
Belanger and Quatrano, 2000b; Nagasato and Motomura,
2002b; Bisgrove and Kropf, 2004). Starting at a
post-telophase stage, when the daughter nuclei have gained
a fully interphase appearance, the possible steps of the
cytokinetic process would be:
(a) Interaction of cytokinetic MTs between each other via
motor proteins, as well as interaction with AFs (and
possibly associated proteins), guides the daughter nuclei
to move apart and/or take their final positions.
(b) Cortical AF cytoskeleton disappears, AF spindle is disorganized and F-actin is gathered in the mid-area
between the daughter nuclei.
(c) A cytokinetic AF disc is formed at the plane that is
determined by the two interdigitating MT systems. In
vacuolated cells, an AF ring is initially organized in the
periphery of the cell, before the formation of the AF
disc. In elongated cells, where the daughter nuclei move
away from each other, a clear zone is left between them.
(d) Vesicles of dictyosome origin and thin, peculiar cisternae of unknown origin, appearing tube-like in sections,
are aligned along this plane, possibly stabilized by
F-actin. This stage probably coincides with stage (c).
(e) Fusion of these vesicles and cisternae results in the
formation of the membranous cytokinetic diaphragm.
(f) The F-actin disc disappears and the daughter cells gain
the interphase actin organization.
(g) Wall deposition from both daughter cells is continued
until the completion of the new cell wall.
CELL WALL MORPHOGENESIS
Microtubule cytoskeleton
The mechanical properties of the cell wall that control its
expansion are mainly determined by the orientation of MFs.
In higher plant cells, cortical MTs have been proven to be
involved in the orientation of MFs, probably by guiding
cellulose synthases to the cell cortex (Williamson, 1991;
Sonobe and Takahashi, 1994; see review in Baskin, 2001;
Smith and Oppenheimer, 2005). Thus, MTs constitute the
main cytoskeletal elements participating in cell wall
morphogenesis of higher plants.
The absence of cortical MTs in brown algal cells suggests
that a different mechanism of cell wall morphogenesis may
operate in them. Experiments using MT inhibitors like
oryzalin did not reveal any effect in the development or
the strength of the cell wall depositing in young zygotes of
Fucales (Kropf, 1992a, b, 1994; Bisgrove and Kropf, 1998).
However, in tip-growing rhizoids of fucalean zygotes, as
well as in apical cells of Sphacelaria, MTs are involved in
cell wall morphogenesis. Their disruption causes abnormal
growth and thickening of the wall of the tip region. It seems
possible that MTs and/or AFs are implicated in the transport
of wall material to the extending wall (Karyophyllis et al.,
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
686
A
B
F I G . 4. Tangential thin sections showing MF orientation in the cell wall of cells of S. rigidula. (A) Control. The difference in the orientation of
MFs between cell wall layers is visible. (B) After cytochalasin treatment. The orientation of the newly deposited MFs is disturbed (arrow). Scale bars:
A = 05 mm; B = 05 mm.
1997; Belanger and Quatrano, 2000b; Karyophyllis, 2003).
Moreover, MTs seem to be implicated in cytokinesis of
S. lomentaria zygotes, by transporting vesicles containing
wall material to the developing diaphragm (Nagasato and
Motomura, 2002b).
Application of antibodies against a-actinin, vitronectrin
and integrin in Sphacelaria, as well as in zygotes of Fucus
has shown that cortical sites involving transmembrane connections between the cytoskeleton and the extracellular
matrix are crucial for the establishment of cell polarity
(Quatrano and Shaw, 1997; Ouichou and Ducreux, 2000;
Brownlee et al., 2001).
Actin filament cytoskeleton
The discovery of a cortical F-actin system in vegetative
cells of brown algae was a strong stimulus for the investigation of the possible role of these AFs in cell wall morphognesis. At first this was checked in the apical cells of
S. rigidula and S. tribuloides (Karyophyllis et al., 2000b).
Thin sections passing through selected regions of the cell
wall (Figs 2C and 4A and B) were examined under TEM.
This wall displays a multilayered structure, similar to that
described very early by Dawes et al. (1960, 1961; see also
Prud’homme van Reine and Star, 1981; Quatrano, 1982;
Tamura et al., 1996). The mature cell wall consists of
four layers, named L1, L2, L3 and L4, going from the
external surface inwards (Fig. 2C). L1 consists of amorphous material and covers all the cell wall externally. L2 is a
thin layer consisting of a few randomly distributed MFs
embedded in an amorphous matrix. L3 is also a thin
layer rich in MFs, which form a ring oriented transversely
to the long axis of the cell. L4 is the innermost and thickest
layer bearing numerous MFs with an axial or fishbone-like
arrangement (Figs 2C and 4A).
It was particularly interesting that the above layers are
gradually depositing along the apical cells, i.e. the cell wall
of the tip area of the cell is very thin and consists only of L1
and L2. L3 starts depositing at the base of the hemispherical
dome of the apex, and L4 at the area where the shape of the
cell becomes cylindrical (Figs 2C and 4A). The abovedescribed MF arrangement coincides with the organization
of AFs in this cell as found after Rh-Ph staining of F-actin
(see above, Fig. 1D).
In order to examine whether the above relationship is a
general phenomenon in brown algal cells, the organization
of MFs and the underlying AFs was investigated by TEM
and Rh-Ph staining, respectively, in a variety of species
and cell types, i.e. subapical and differentiating cells of
S. rigidula, apical cells of D. dichotoma, subapical cells
of Choristo-carpus tenellus and meristematic cells of
D. dichotoma. In all cases a cortical AF system was present,
always oriented parallel to the depositing MFs of the innermost wall layer (Karyophyllis et al., 2000b; Katsaros et al.,
2002). This coincidence suggests a possible involvement of
AFs in the orientation of MFs.
To confirm the above hypothesis, F-actin was disrupted
with CB, in order to ascertain whether MF deposition would
be affected. Treatment of S. rigidula and D. dichotoma thalli
with 100 mg mL1 CB for 24–36 h caused a gradual disruption of AFs. TEM examination of treated material
showed that the MFs of the internal wall surface did not
exhibit the clear parallel arrangement observed in normal
cells. Instead, a rather random orientation of loose and thin
MFs was found (Fig. 4B). It must be noted that the cell wall
of fully differentiated cells was normal. Cytochalasin affected only cells where wall deposition was in progress during
the treatment (Karyophyllis et al., 2000b; Katsaros et al.,
2002). A similar suggestion has been made by Mizuta et al.
(1994), who reported that the helicoidal pattern of MF orientation in the giant green alga Boergensenia was disrupted
after cytochalasin treatment.
Although the coincidence of AF–MF orientation has
not been confirmed until now by direct observation under
TEM, the mutual alignment between cortical AFs and MFs
is strong evidence that AFs are involved in cell wall
morphogenesis of brown algae. This means that a different
mechanism of cell wall morphogenesis is functioning in this
group. In this mechanism, AFs seem to be involved,
playing a role similar to that of cortical MTs in higher
plant cells.
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
CENTROSOME
Structure: centrosomal proteins
Vegetative cells of brown algae bear detectable centrosomes during their whole cycle; in this they differ from
higher plants and resemble animal cells. Centrosomes in
brown algae were found very early by light microscopy
(Strasburger, 1897; Swingle, 1897; Mottier, 1900;
Yamanouchi, 1909), and described in more detail by
electron microscopy (Berkaloff, 1963; see also Manton
and Clarke, 1950; Manton, 1957). Each centrosome consists
of two centrioles that lie at about right angles to one another
and in close proximity at one end (proximal). In animal
cells, pericentriolar material (PCM) surrounds the centrioles
and is in part organized by them (Bobinnec et al., 1998).
PCM is the site of MT nucleation (Wheatley, 1982;
Balczon, 1996; Doxsey, 2001) through g-tubulin containing
complexes (Moritz and Agard, 2001), although other
proteins also appear to be involved in this process. The
centrosome comprises hundreds of proteins, several of
them serving as docking sites for a number of regulatory
and other activities (Doxsey et al., 2005).
The Ca2+-modulated contractile protein centrin was, at
first, found associated with basal bodies of green flagellates
(Salisbury et al., 1984; Huang et al., 1988a, b). After its first
localization and identification, centrin was found in a great
variety of eukaryotic cells, in relation with basal bodies,
mitotic spindles, spindle poles, centrosomes, etc. In higher
plant cells centrins or their homologues have been found in
the cell plate, on the nuclear surface and in the spindle poles
(Wick and Cho, 1988; Del Vecchio et al., 1997; StoppinMellet et al., 1999, 2000). In cells bearing centrosomes,
centrin can be used as a centrosome marker (Paoletti
et al., 1996).
In brown algae, centrin was detected in the basal apparatus of motile male gametes (Melkonian et al., 1992;
Katsaros et al., 1993). It was localized in protein structures
connecting the basal bodies to each other or to the nucleus
(nucleus–basal body connector) (Katsaros et al., 1993). This
connection is quite strong and extraction of motile cells
using detergents does not separate basal bodies from the
nuclei. Centrin was also localized in developing gametangia, zygotes and vegetative cells of brown algae (Katsaros
et al., 1991; Katsaros and Galatis, 1992; Karyophyllis,
1995; Bisgrove et al., 1997; Nagasato et al., 1998,
1999a, 2000). In vegetative cells it was always associated
with the centrosome site during the whole cell cycle. The
centrin-containing structure in centrosomes of vegetative
cells has not been identified. However, considering
the close relationship between centrosomes and nuclei, it
could be speculated that centrin has a similar connecting
role. Studies in ‘lower’ eukaryotes and, more recently, in
animal cells have established that centrin-containing fibres
play a role in the dynamic behaviour of centrosomes,
through control of the position and orientation of centrosomal structures, and also in the control of centriole duplication (Baum et al., 1988; Marshall et al., 2001; Salisbury
et al., 2002; Kilmartin, 2003; Salisbury, 2004). Recently
cDNA and the genomic DNA of the centrin gene of
687
S. lomentaria was isolated and analysed. The protein
deduced exhibited 84 % homology to the Chlamydomonas
centrin (Nagasato et al., 2004).
Another centrosomal protein, g-tubulin, has been found
in various cell types of animals, plants and fungi, associated
with spindle poles or other MTOCs, as well as along MTs
(Oakley and Oakley, 1989; Oakley et al., 1990; Stearns
et al., 1991; Liu et al., 1995; Zheng et al., 1995; Joshi
and Palevitz, 1996; Vaughn and Harper, 1998; Panteris
et al., 2000) and it is generally accepted that it participates
in MT nucleation (Moritz et al., 1995; Oakley, 2000). By
using a special antibody, g-tubulin was always found in
brown algal cells at the centrosome area (Karyophyllis,
2003; Karyophyllis et al., 2005b). The pattern of g-tubulin
localization in S. rigidula coincides with that in animal cells
and fungi (Pereira and Schiebel, 1997; Oakley, 2000) supporting its nucleating role (Karyophyllis et al., 2005b). The
intensity of the immunofluorescence reaction of g-tubulin
changes during mitosis, with a maximum at metaphase. This
is probably caused by an increase in g-tubulin in the spindle
pole due to the accumulation of the protein that is dispersed
in the cytoplasm. Similar accumulation of g-tubulin has
been observed in animal cells during mitosis (Vorobjev
et al., 2000). As the centrosomes ‘mature’ they recruit additional g-tubulin ring complexes and several other proteins,
resulting in an increase in the nucleation capacity of the
centrosome (reviewed by Blagden and Glover, 2003). This
possibly reflects the increased number of MTs nucleated at
mitosis compared with interphase (Kuriyama and Borisy,
1981), which is the case in brown algae like D. dichotoma,
as it is revealed with electron microscope observations
(Katsaros and Galatis, 1992).
The identification in brown algae of complexes containing g-tubulin or/and its precursors is an interesting task from
an evolutionary perspective, since the heterocont group
diverged early in eukaryotic evolution (Baldauf, 2003).
These structures are present not only in animals and
fungi (Zheng et al., 1995; Jeng and Stearns, 1999; Wiese
and Zheng 1999; Schiebel, 2000; Moritz and Agard, 2001),
but also in plants (Stoppin-Mellet et al., 2000; Dryková
et al., 2003).
Centrosome cycle
During interphase, brown algal centrosomes are located
close to the nucleus, sometimes in a shallow depression of
the nuclear envelope (Bouck, 1965; Neushul and Dahl,
1972; Katsaros, 1980; Katsaros et al., 1983; Motomura,
1991, 1994; Katsaros and Galatis, 1992), MTs radiate out
of the pericentriolar material, some of them surround the
nucleus, while others traverse the cytoplasm towards the
cell cortex.
At prophase the activity of the two oppositely located
centrosomes appears to be increasing, and numerous
short MTs or MT bundles are assembled, forming asterlike configurations. At the same time the depressions of
the nuclear envelope become broader and the MTs
seem to contact the nuclear envelope, interacting with an
688
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
electron-dense material that appears at its cytoplasmic face.
This interaction probably causes the breakage of the already
weakened nuclear envelope at the poles. After the formation
of the polar gaps, spindle is formed by the extension of MTs
from the centrosomes towards the chromosomes.
Duplication of centrosomes usually occurs during the
S phase of the cell cycle, and the daughter centrosomes
separate and migrate to opposite sides of the nucleus
(Kuriyama and Borisy, 1981). In most cases of brown
algal cells examined so far, two oppositely placed centrosomes (two centriole pairs) are visible during most of
interphase, since duplication and separation of centrosomes
occurs soon after cytokinesis, and thus they are in place
before any sign of the onset of mitosis (Katsaros and
Galatis, 1992; Rusig et al., 1994). MTs seem to be involved
in the separation and migration of the daughter centrosomes
to the poles. Depolymerization of MTs after Nz treatment in
S. rigidula did not affect centrosome duplication, but disturbed their separation. Prolonged treatment caused multicentrosomal nuclei (up to four in one nucleus). This means
that MT disruption inhibits cell division, but not the centrosome cycle, that proceeds independently (Karyophyllis
et al., 1997; Karyophyllis, 2003). This observation is in
agreement with the idea that the centrosome is not under
the cell cycle control, rather it exerts control over the cell
cycle (Doxsey et al., 2005). However, since anti-centrin
immunofluorescence was used for centrosome identification, the possibility that the centrin spots represent separated
centrioles or PCM fragments and not complete centrosomes
should not be excluded (Karyophyllis et al., 1997). Interestingly, centrosome separation was recently found associated with Golgi apparatus. Unexpectedly, disorganization of
Golgi after treatment with brefeldin A seems to inhibit
centrosome movement
to the
opposite
poles
(V. Varvarigos et al., unpublished).
Apart from centrosome separation, MT dynamics seem to
play a significant role in the structure and function of centrosomes in dividing cells of brown algae. In apical cells of
S. rigidula, after Nz treatment, centrin was localized at
cytoplasmic sites far from the centrosomes. It was suggested
that disruption of MTs probably caused fragmentation of
centrosomes or dispersal of centrosomal material, or even
shift of centrosomes from their correct position
(Karyophyllis et al., 1997; Karyophyllis, 2003). In cells
recovering from Nz treatment, a pair of long, parallel,
rod-like structures, showing positive centrin reaction
appeared at the centrosome sites (Karyophyllis et al.,
1997; Karyophyllis, 2003). These structures probably
represent abnormally differentiated centrioles, similar to
‘primary cilia’ described by Wheatley et al. (1996). A similar phenomenon was described by Alieva and Vorobjef
(1991) in cultured cells of pig embryo. These authors
observed quite elongated centrioles (1.5–5 times longer
than the usual).
Recent studies, using polyspermic zygotes of the brown
alga S. lomentaria, have shown that in brown algae the
cytokinetic plane is determined by the position of the
centrosomes after mitosis (Nagasato et al., 1999b;
Nagasato and Motomura, 2002a) and not by the
spindle as it was proposed earlier (Bisgrove and Kropf,
2001). Similar results have been reached by Bisgrove
et al. (2003) in asymmetric division of fucoid zygotes,
where it was suggested that the division plane is determined
by the telophase nuclei.
CONCLUDING REMARKS: FUTURE
PERSPECTIVES
The study of both fine structure and cytoskeleton organization shows that the cytoskeletal elements studied so far are
implicated in brown algal cell morphogenesis. The novelty
is the role of AFs and the probable co-operation between
them and MTs. The dominating role of cytoskeleton, and
especially AFs, in morphogenesis of brown algal cells is
expressed in the following processes:
(a) The structural and functional interconnection between
the cytoskeleton, the plasma membrane and the extracellular matrix (= cell wall). This is analogous to the
‘cell periphery complex’ proposed for higher plant cells
(Baluska et al., 2000) in which cortical MTs are
involved. In both model systems of brown algae studied,
namely zygotes and vegetative cells, a cortical AF cytoskeleton seems to play a critical role. This scheme has
been considered as a more ‘primitive’ complex that has
been replaced by the cortical MTs in higher plant cells
(Baluska et al., 2000, 2001).
(b) Polarization. The implication of radial F-actin configurations in cell polarization seems to be a general
phenomenon in brown algae, as it is in a great variety
of other organisms.
(c) Organization of the mitotic spindle. Co-operation or at
least coexistence occurs between spindle MTs and AFs
during mitosis. The inhibition of anaphase after
treatment with CB indicates a direct or indirect role
of F-actin in mitosis of brown algal cells.
(d) Cytokinesis. Fluorescence microscopy of both normal
and experimentally treated material has revealed an
involvement of F-actin in the cytokinetic process.
This finding, combined with the absence of any of
the known cytokinetic systems (phragmoplast, phycoplast or cleavage furrow) and the presence of centrosomes, showed that a particular cytokinetic mechanism
operates in brown algal cells.
All the above need further investigation. For the first, the
mechanism by which the ‘cell periphery complex’ regulates
the deposition and orientation of the nascent MFs remains to
be clarified. Several studies using freeze fracture have
shown putative cellulose synthase complexes (TCs) on
the plasma membrane of brown algal cells (Peng and
Jaffe, 1976; Katsaros et al., 1996; Reiss et al., 1996;
Tamura et al., 1996; Schüssler et al., 2003). Similar
approaches, combined with rapid freeze-fixation, could possibly give information on the relationship between the cortical F-actin cytoskeleton and the transmembrane synthase
complexes. An analogous relationship between cortical
MTs and rosette-type TCs has been suggested for cellulose
synthesis in higher plants (Mizuta, 1992; Sonobe and
Takahashi, 1994).
Katsaros et al. — Cytoskeleton and Morphogenesis in Brown Algae
The role of F-actin and the centrosome in the cytokinetic
process is also a very stimulating issue that needs further
study. Together with it, the investigation of the mechanism
determining the cell division plane is also a rather attractive
target.
ACKNOWLEDGEMENTS
This study was partially supported by grants from the
University of Athens and from the Hellenic Ministry of
National Education and Religious Affairs and the EU, in
the frames of the Operational Programme for Education and
Initial Vocational Training (O.P. ‘Education’, project
‘Pythagoras’). The authors would like to thank
Mr V. Varvarigos for his assistance in making the drawings.
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