• brown alga;
  • defense;
  • development;
  • Ectocarpus;
  • metabolism;
  • phaeophyceae;
  • pheromone;
  • reproduction


  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References


II.Diversity and taxonomy, distribution and ecology321
V.Interactions with the environment327


Brown algae share several important features with land plants, such as their photoautotrophic nature and their cellulose-containing wall, but the two groups are distantly related from an evolutionary point of view. The heterokont phylum, to which the brown algae belong, is a eukaryotic crown group that is phylogenetically distinct not only from the green lineage, but also from the red algae and the opisthokont phylum (fungi and animals). As a result of this independent evolutionary history, the brown algae exhibit many novel features and, moreover, have evolved complex multicellular development independently of the other major groups already mentioned. In 2004, a consortium of laboratories, including the Station Biologique in Roscoff and Genoscope, initiated a project to sequence the genome of Ectocarpus siliculosus, a small filamentous brown alga that is found in temperate, coastal environments throughout the globe. The E. siliculosus genome, which is currently being annotated, is expected to be the first completely characterized genome of a multicellular alga. In this review we look back over two centuries of work on this brown alga and highlight the advances that have led to the choice of E. siliculosus as a genomic and genetic model organism for the brown algae.

I. Introduction

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

The brown algae belong to the division Heterokonta and are therefore only very distantly related to the three most intensely studied eukaryotic groups, the animals, fungi and green plants (Baldauf, 2003; Davis, 2004; Fig. 1a). This independent evolutionary history has furnished brown algae with many novel metabolic, physiological, cellular and ecological characteristics, including a complex halogen metabolism, cell walls containing many unusual polysaccharides and high resistance to osmotic stress. Developmental processes are particularly interesting in this group, which evolved complex multicellularity independently of the three other aforementioned major groups. From a more applied point of view, the evolutionary history of the brown algae also underlies the high commercial value of several members of the group in the sense that they have evolved novel biomolecules such as polysaccharides and defense elicitors that have a wide range of applications in industry (Klarzynski et al., 2000; McHugh, 2003).


Figure 1. Phylogeny of brown algae and Ectocarpales. (a) Position of brown algae within the eukaryotes (adapted from Baldauf, 2003). Brown algae belong to the heterokont phylum, which is phylogenetically distant from land plants and the green and red algae. Photosynthetic organisms are framed. (b) Position of the Ectocarpales (in bold) within the brown algae (adapted from Kawai et al., 2007).

Download figure to PowerPoint

It is important to note, however, that whilst the independent evolutionary history of the brown algae is the source of much of the interest of this group, it can also be seen as a handicap because the well developed model organisms from the plant and animal lineages are of limited relevance to brown algal biology. Specialized brown algal models have been developed in specific domains, for example members of the fucoids for cell biology approaches (see references in Corellou et al., 2005), but a polyvalent model organism that allows access to a wide range of questions at the molecular level has been lacking. This situation is changing with the development of genomic and genetic tools for the filamentous brown alga Ectocarpus siliculosus. The sequencing of the genome of this alga has recently been completed and the sequence is currently being annotated ( It is therefore an opportune moment to look back at the emergence of Ectocarpus as a model organism.

Research on Ectocarpus began in the 19th century with descriptions of species and taxonomy, followed by studies aimed at unravelling reproduction and life history. Other major aspects that have been studied include the sexual pheromones and infection of Ectocarpus by viruses. Research has also been carried out on ultrastructure, photosynthesis and carbon uptake, gamete recognition and resistance to anti-fouling agents. Several eukaryotic parasites of Ectocarpus have been described. A proposition to adopt Ectocarpus as a general model organism for the brown algae was made in 2004 (Peters et al., 2004a). This proposition was based partly on this alga's long history as an experimental organism, but also took into account several features that make Ectocarpus an interesting model for genetic and genomic approaches. These features include its small size, the fact that the entire life cycle can be completed in Petri dishes in the laboratory (Müller et al., 1998), its high fertility and rapid growth (the life cycle can be completed in 3 months), the ease with which genetic crosses can be carried out and the relatively small size of the genome (200 Mbp compared with 1095 and 640 Mbp for Fucus serratus and Laminaria digitata, respectively; Le Gall et al., 1993; Peters et al., 2004a).

Here we present an overview of the work that has been carried out on Ectocarpus over the last two centuries and discuss how the availability of a number of genomic tools, in particular the complete genome sequence, is expected to accelerate research in many domains of brown algal biology in the coming years.

II. Diversity and taxonomy, distribution and ecology

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

1. Diversity and taxonomy

Dillwyn (1809) published the first valid description of Ectocarpus (using the name Conferva siliculosa) based on material collected by W. J. Hooker on ‘rocks in the sea at Cromer and Hastings’. These English localities lie in Norfolk and East Sussex, respectively. Type material, collected by Hooker in 1807, is housed at BM (BM000685585 and BM000685588) under the name C. confervoides. Lyngbye (1819) described the genus Ectocarpus based on material from Denmark and cited C. siliculosa Dillwyn as basionym. The correct nomenclature, therefore, is E. siliculosus (Dillwyn) Lyngbye (see Silva et al., 1996 for further details). Ectocarpus siliculosus is the type species of the order Ectocarpales, which includes most of the smaller brown algae. Originally regarded as phylogenetically primitive, molecular systematics has shown the Ectocarpales to belong to a group of brown algal orders that evolved quite recently. They are closely related to the large and highly differentiated Laminariales, which are major components of coastal marine floras (Rousseau & de Reviers, 1999; Draisma et al., 2003; Cho et al., 2004; Kawai et al., 2007, Fig. 1b). Many species have been described in Ectocarpus (, April 2007, listed 392 taxa of Ectocarpus, of which 98 are flagged as ‘current’; numerous strains are publicly available at the Culture Collection of Algae and Protozoa (CCAP) in Oban, UK; Gachon et al., in press). However, only E. fasciculatus Harvey (1841) is currently recognized as a second, well defined species, based on morphology (Russell, 1966, 1967a), crossing studies and sequence analyses (Stache-Crain et al., 1997). Crossing experiments have shown that the taxon E. siliculosus may represent a species complex (Stache-Crain et al., 1997) and ongoing, refined analyses are expected to resolve this complex, increasing the number of recognized species. Identification of different species of Ectocarpus based on morphology is difficult because of the plasticity of the commonly examined features (habit, branching pattern, size of sporangia). In addition, the two generations of a species may differ considerably (Müller, 1972a; Kornmann & Sahling, 1977).

2. Distribution

Ectocarpus siliculosus is distributed worldwide in temperate regions, but does not occur in the tropics and south of the Antarctic convergence (Stache, 1990; Wiencke & Clayton, 2002). It occurs in fully marine and in low-salinity habitats (e.g. five practical salinity units (psu) in Finland) and has even been recorded at a freshwater site in Australia (West & Kraft, 1996) and in a salt-polluted river in Germany (Geissler, 1983). Records of E. fasciculatus are mainly from the North Atlantic but there are some from Korea, Chile, South Georgia and South Africa (, 2007). On the shore, Ectocarpus occurs from high intertidal pools to the sublittoral. It is found on abiotic substrata (rocks, wood, plastic, ship hulls) and epiphytic on macrophytes or free-floating (Russell, 1967a,b, 1983a,b). As a result of its ability to grow on a range of abiotic substrates, Ectocarpus is a common fouling alga (Morris & Russell, 1974).

3. Ecology

There have been a limited number of ecophysiological and ecological studies on Ectocarpus. Growth rate is dependent on temperature, and there is evidence that temperature also influences the life cycle, at least in some strains (Müller, 1963; Bolton, 1983; see Section III.2: Sporophyte and gametophyte architecture). The thermosensitivity of different strains suggests that there is genetic heterogeneity within the Ectocarpus genus (Bolton, 1983). A similar result was obtained for osmo-acclimation (Thomas & Kirst, 1991a,b; see Section V.2: Abiotic stresses). In the field, Ectocarpus is a short-lived annual which may dominate the ectocarpoid flora on kelps (Russell, 1983a,b). Despite the commonness of Ectocarpus, there are few data concerning phenology in the field (mostly to be extracted from floristic works) and nothing precise on seasonality or habitats of the two generations.

The availability of the E. siliculosus genome sequence is expected to facilitate the analysis of the ecology of this species by providing a basis for the development of molecular markers. Particularly important challenges in this respect will include the identification of the sex locus and of genes specifically expressed in the two generations, as these will provide molecular tools that can be used to investigate several aspects of the life cycle under field conditions. Molecular markers will also allow the exploration of genetic polymorphism among Ectocarpus species from multiple locations across the globe.

III. Development

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

1. Life cycle and reproduction

Male and female gametes are morphologically identical in Ectocarpus (isogamy) but differ with respect to their physiology and their behaviour: female gametes settle sooner and produce a pheromone whilst male gametes swim for longer and are attracted to the pheromone produced by the female. Studies on the reproduction of Ectocarpus began with the observation of sexual fusions involving the attraction of male gametes to settled female gametes from field thalli of E. siliculosus in Naples, Italy (Berthold, 1881). These findings were hotly debated until Sauvageau (1896, 1897) and Oltmanns (1899) succeeded in repeating the experiment. Gamete fusions in Ectocarpus were later used by Hartmann (1934) to support his erroneous theory of relative sexuality (see Müller, 1976a for details). Knight (1929) identified the young, unilocular sporangium as the site of meiosis, Papenfuss (1935) and Kornmann (1956) published major contributions on the life history of Ectocarpus, and Boalch (1961) developed refined culture techniques. The entire life history of E. siliculosus from Naples was finally unravelled by Müller (1964, 1966, 1967, 1972b) using clonal cultures and chromosome counts. It is schematized in Fig. 2. The basic life history of E. siliculosus involves an alternation between the sporophyte and dioecious gametophytes, and sex determination is genotypic (Müller, 1967). Male and female gametophytes are morphologically indistinguishable. One of the problems with understanding the life history was that sporophytes and gametophytes are difficult to distinguish morphologically. Another problem was that zoids from plurilocular reproductive organs have different functions according to the generation forming them: on sporophytes they contain asexual zoospores that directly reproduce the sporophyte, while on gametophytes they contain gametes. Further complications include the parthenogenesis of unfused gametes, which develop into haploid parthenogenetic sporophytes morphologically indistinguishable from diploid sporophytes, and heteroblasty (different fates) of spores from unilocular sporangia, developing either into gametophytes or into sporophytes. Furthermore, life cycle generation is not determined rigidly by ploidy (Müller, 1967).


Figure 2. Life cycle of Ectocarpus siliculosus. Diploid sporophytes produce meiospores (by meiosis) in unilocular sporangia (UL). Meiospores grow into male or female gametophytes (dioecism). Gametophytes produce gametes in plurilocular gametangia (PL). Fusion of gametes produces a zygote that grows into a diploid sporophyte, completing the sexual cycle. Unfused gametes may grow parthenogenetically and form a parthenosporophyte, which is indistinguishable from the diploid sporophyte. Both sporophytes and parthenosporophytes can reproduce themselves asexually by the production of mitospores in plurilocular sporangia.

Download figure to PowerPoint

An important challenge for the future will be the characterization of the genetic mechanisms that control life cycle progression in Ectocarpus. This will require the development of methodologies for positional cloning of mutated loci and of genome-wide methods to analyse gene expression throughout the life cycle. Work is currently ongoing in several groups to develop these techniques.

2. Sporophyte and gametophyte architecture

Ectocarpus siliculosus is a small filamentous alga that grows to c. 30 cm in length in nature, but it may become fertile in the laboratory at 1–3 cm.

Sporophyte development is initiated with the germination of the zygote. The first division produces two cells of identical developmental fate (Peters et al., 2004b). Subsequent mitoses lead to the formation of a basal (or prostrate) filamentous structure, defining the early sporophyte (Fig. 3a). Phaeophycean hairs, that is, hyaline filaments devoid of plastids developing from a basal meristem, are absent in Ectocarpus but present in the sister genus Kuckuckia. However, in Ectocarpus the distal end of filaments, or of plurilocular sporangia, may be less pigmented and resemble a hair; such structures may be referred to as pseudo-hairs (Cardinal, 1964; Pedersén, 1989). If the growth conditions are favourable (Ravanko, 1970), erect filaments (called ‘upright’ filaments) emerge after a few days, contributing to the establishment of an overall filamentous architecture (Fig. 3c).


Figure 3. Morphology of Ectocarpus siliculosus. Photographs of 1-wk-old vegetative sporophyte (a) and gametophyte (b) and schemes representing the whole body of the mature sporophyte (c) and gametophyte (d) after 6 wk of growth. (e) Plurilocular sporangium and gametangium (occurring on the sporophyte and the gametophyte, respectively) before (left) and after (right) release of zoids. (f) Unilocular sporangium from sporophyte. Sporangia and gametangia can be either sessile or pedicellate (Kim & Lee, 1992).

Download figure to PowerPoint

The typical structure of a vegetative cell is illustrated in Fig. 4. Features common to all brown algal cells include a chloroplast surrounded by four membranes, arranged as two double-membraned envelopes (chloroplast envelope, CE). The second envelope is loosely associated with the chloroplast and forms part of the chloroplast endoplasmic reticulum (CER). The lamellae of the chloroplast are composed of three thylakoids, which are absent from the pyrenoid space (Bouck, 1965; Oliveira & Bisalputra, 1973). In Ectocarpus the chloroplast is ribbon-shaped. The size and number of chloroplasts may vary within the same organism (Ravanko, 1970). Other typical features of Ectocarpus include several prominent and pedunculated pyrenoids on the inner face of the chloroplast, which are used as a taxonomic marker for the Ectocarpales (Evans, 1966; Rousseau & de Reviers, 1999). Chloroplast endoplasmic reticulum and CE envelopes also surround pyrenoids, this time being tightly adjacent. A third external envelope, called the pyrenoid sac, surrounds the pyrenoid but has no connection with the reticulum system (Bouck, 1965). The nuclear envelope is continuous with the CER, which is itself in close vicinity to the Golgi apparatus (Bouck, 1965; Oliveira & Bisalputra, 1973). It has been hypothesized that these connections create a complex network of membranes allowing photosynthates to be efficiently transferred from the chloroplast to the Golgi apparatus, the latter being also in direct contact with the CER (Oliveira & Bisalputra, 1973). Cytoplasmic ER is dispersed throughout the cytoplasm and is mainly rough (Oliveira & Bisalputra, 1973). Osmiophilic bodies (OSB), which are thought to contain lipids, are dispersed throughout the cytoplasm and probably originate from the CER (Oliveira & Bisalputra, 1973). They have been observed within the cell wall and also external to it. Vacuoles can be either large structures occupying peripheral locations (Oliveira & Bisalputra, 1973) in fixed material, or most of the cellular space (as in land plant cells; Knight, 1929, confirmed by data from our laboratory, after staining with cresyl blue or neutral red, on sporophytic filaments). The nuclear region encompasses two centrioles, which are considered as a microtubule organizing centre (MTOC; Katsaros et al., 1991). The chromosome number in the haploid nucleus is estimated to be c. 25 (Peters et al., 2004a). Mitochondria are preferentially peripherally located. They are maternally inherited (Peters et al., 2004b). The cell wall consists of a fibrillar matrix (see Section IV.1: Photosynthesis and carbohydrate metabolism) with several plasmodesmata distributed uniformly along the cross walls. Upon ageing, cell wall ingrowths occur, accompanied by a reduction in the size of the nucleus, mitochondria, ER and Golgi, followed by the disintegration of the chloroplasts and finally by autolysis of the cytoplasm (Oliveira & Bisalputra, 1977a,b).


Figure 4. General ultrastructure of a vegetative cell of Ectocarpus siliculosus. The general ultrastructure of a vegetative cell is similar in both prostrate and erect filaments (Oliveira & Bisalputra, 1973). The different compartments of the cell are illustrated (see text for details). Lines represent membranes and define subcellular compartments, except for thylakoids, drawn as a thick black line. Depending on their type and age, vegetative cell size varies from 10 to 35 µm in length, and 5 to 15 µm in width (under laboratory culture conditions).

Download figure to PowerPoint

Most of the reproductive organs are carried by upright filaments (Müller, 1964). Two types of reproductive organs are produced by the sporophyte: plurilocular and unilocular sporangia. Plurilocular sporangia are cone-shaped three-dimensional structures of variable size (Knight, 1929), composed of a large number of locules with different shapes (Baker & Evans, 1973b; Fig. 3e). These locules are generated by several successive mitoses and each gives rise to a single zoospore, which is released through the apex of the sporangium. The mitospores, which are biflagellate and competent for swimming shortly after their release (Müller, 1980), are a means of vegetative reproduction. Unilocular sporangia are born on the sides of branches (Fig. 3f, Baker & Evans, 1973a). A single meiosis occurs within the single thick-walled locule (Baker & Evans, 1973a) and this is followed by several mitoses, which generate about a hundred meiospores, half of which are female and half male (Müller, 1980). Müller (1963) and Ravanko (1970) reported that plurilocular sporangia were produced when the external temperature was relatively high (c. 20°C or summer), whereas unilocular sporangia were produced when the temperature was lowered to 13°C (mimicking winter conditions). However, this temperature dependence is not observed in many strains of Ectocarpus (A. F. Peters, unpublished data). Meiospores germinate to produce haploid, dioecious gametophytes (Fig. 3b,d). These are filamentous organisms similar to the sporophyte, but with two important differences. Firstly, the meiospore germinates asymmetrically to produce a rhizoid and an upright filament, so no prostrate structure forms (A. F. Peters, unpublished data). Secondly, the thallus is more ramified than that of the sporophyte (Müller, 1980). Gametophytes produce only plurilocular gametangia and these are similar structurally to plurilocular sporangia on sporophytes. Gametes resemble mitospores in terms of their size and motility.

The developmental patterning varies greatly across the Ectocarpus species complex and is also dependent on the environment, growth conditions and even the age of the algae for some features (Ravanko, 1970). This plasticity is observed for the branching frequency and the number, shape, structure and positioning of the reproductive organs on filaments (Knight, 1929; Müller, 1980; Kim & Lee, 1992). Phytohormones, especially cytokinins, have also been reported to influence the development of Ectocarpus sporophytes (Pedersén, 1968, 1973).

As already mentioned, brown algae are interesting for developmental studies because they have independently evolved complex multicellularity. Ectocarpus is a relevant model to address this problem as it is closely related to complex algae such as the Laminariales and the Fucales. However, developmental processes in Ectocarpus are clearly simplified compared with its morphogenetically more complex sister families, which provides an advantage for the detailed dissection of these developmental processes. In particular, the simple growth pattern of the uniseriate, branched filaments represents an ideal system for the combined application of genetic and mathematical modelling approaches to understanding developmental patterning and then addressing the issue of the evolution of development and multicellularity.

3. Gametes and spores

Several electron microscopy studies of the motile cells (zoids) of Ectocarpus have been reported (Baker & Evans, 1973a,b; Lofthouse & Capon, 1975; Maier, 1997a,b). The typical structure of an Ectocarpus zoid is illustrated in Fig. 5.


Figure 5. General intracellular structure of Ectocarpus siliculosus zoids. A scheme representing the different compartments of an Ectocarpus siliculosus zoid cell from a plurilocular sporangium or gametangium is presented (see text for details). Lines represent membranes and define subcellular compartments, except for thylakoids, drawn as a thick black line.

Download figure to PowerPoint

Ectocarpus zoids correspond to the ‘primitive’ type of brown algal zoid according to Kawai (1992). Gametes and spores typically contain a single chloroplast with a pyrenoid (Baker & Evans, 1973a; Lofthouse & Capon, 1975; Maier, 1997a). As in the vegetative cells, lamella are composed of three thylakoids, and the nuclear envelope is in continuity with the chloroplast endoplasmic reticulum (Maier, 1997a). The nucleus of the male gamete is rich in heterochromatin. Several dictyosomes are present in gametes and spores (Baker & Evans, 1973b; Maier, 1997a). The Golgi apparatus of mitospores is very active both before and after release (Baker & Evans, 1973b). This secretory activity may have important functions in the biosynthesis of the adhesive required for gamete adhesion and for the synthesis of new cell wall compounds during germination.

Ectocarpus gametes and spores are characterized by two flagella with lateral insertion. One is oriented forward and equipped with mastigonemes (hairs), and propels the cell with meandering beats. The second is oriented obliquely backwards and has no mastigonemes. Most of the time this posterior flagella is passively dragged, but occasional lateral beats induce abrupt changes in direction of up to 180° (Geller & Müller, 1981). Gametes and spores have a concave depression at the level of the eyespot into which the swelling of the proximal part of the posterior flagellum fits (Baker & Evans, 1973b; Kreimer et al., 1991; Maier, 1997a). The possible function of the eyespot is to reflect and focus incident light on to the site of photoreception (Kawai et al., 1990; Kreimer et al., 1991). Zoids are capable of positive phototaxis and their posterior flagellum shows strong autofluorescence when irradiated by blue light (450 nm). The photoreceptor pigment is a flavoprotein, which is periodically shaded by a carotenoid stigma (Müller et al., 1987; Kawai, 1988). The acronema (the whiplash tip) is extremely sensitive to mechanical stress and plays an important role in establishing the initial sexual contact between gametes. The details of the flagellar apparatus of both female and male gametes have been studied by electron microscopy (Müller & Falk, 1973; Maier, 1997b). Despite their different behaviours, no difference in their fine structure has been detected (Müller & Falk, 1973).

The pheromone that attracts male gametes to female gametes is an unsaturated hydrocarbon. The substance initially identified was ectocarpene (all-cis-1-(cycloheptadiene-2′,5′-yl)-butene-1) (Müller et al., 1971; Müller, 1976b, 1978; Müller & Schmid, 1988) but more recently Boland et al. (1995) have shown that a thermally labile cyclopropyl precursor, pre-ectocarpene, is more active by three orders of magnitude and is thus the actual pheromone.

There is evidence that cell-to-cell recognition between Ectocarpus gametes is mediated by N-acetyl glucosamine residues exposed on the plasma membrane of the female gametes and that these residues are specifically recognized by a receptor on the male gamete (Schmid, 1993; Schmid et al., 1994a). In addition, the lectins concanavalin A (Con A) and Aleuria aurantia agglutinin (AAA) bind specifically to the anterior flagella of Ectocarpus gametes and the molecules with which they interact also may be involved in the gamete recognition process (Maier & Schmid, 1995).

Zoids from plurilocular and unilocular Ectocarpus sporangia share similar overall intracellular structures, but there are some important differences: plurilocular zoids are smaller and swim faster with more rapid changes of direction (Baker & Evans, 1973a). Zoids from unilocular sporangia are strikingly different from the other Ectocarpus cell types (both plurilocular zoids and vegetative cells) in that the nucleus is physically separated from the chloroplast. Moreover, secretory activity is lower than in zoids from plurilocular sporangia (Baker & Evans, 1973b).

Gametes of Ectocarpus have also been used to study chemotaxis (Boland et al., 1983), receptor modelling (Boland et al., 1989) and phototaxis (Kawai et al., 1990; Kreimer et al., 1991). Protocols for isolation and biochemical characterization of plasma membrane and CER membrane have also been developed (Schmid et al., 1992). Together with more recently developed molecular tools, these methods now offer access to a range of interesting biochemical events that take place during gamete interaction at fertilization, such as chemoreception, cell-cell recognition and fusion processes.

IV. Metabolism

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

1. Photosynthesis and carbohydrate metabolism

Marine environment constrains several aspects of photosynthesis in brown algae. First, carbon dioxide is not the main source of inorganic carbon (Ci). Indeed, seawater in equilibrium with air contains only 13 µM CO2, but ~ 2 mM anionic carbon, mainly in the form of inline image (Beer, 1994). Secondly, when seaweeds are immersed they do not receive the full light spectrum. Light absorption increases with water depth and varies according to the wavelength: red light (650 nm) is absorbed first, followed by purple (400 nm) and yellow (550 nm) light. By contrast, green (500 nm) and blue (450 nm) light display a strong penetration: at 10 m depth, red light is almost fully absorbed whereas the absorption of green and blue light is not significant.

Adapted to these conditions, brown algae differ from land plants by the pigment composition of their light-harvesting complexes (LHCs). Chlorophyll c (chlc) and the carotenoid fucoxanthin are indeed the main light-haversting pigments of brown algae. Their presence broadens the absorption spectrum toward the green light relative to the chlorophytes. Brown algae possess two different LHCs associated with photosystems I and II (PSI and PSII), but their pigment composition is controversial. Barrett & Anderson (1980) described a fucoxanthin chla/c protein and a violaxanthin-enriched chla/c protein. Conversely, Alberte et al. (1981) reported a chla/c protein devoid of fucoxanthin and a second LHC containing chla and fucoxanthin but not chlc. More recent analyses showed that the LHCs associated with PSI and PSII are in fact virtually identical with respect to their pigmentation and peptide composition. All complexes bound chla, chlc and fucoxanthin in the proportion 6 : 2 : 7, but the LHC associated with PSI is significantly enriched in violaxanthin (De Martino et al., 2000). As in plants, these accessory pigments transfer energy to chla within the photosynthetic reaction centres (Grossman et al., 1995). Violaxanthin does not participate in light-harvesting but is involved in photoprotection (Demmig-Adams & Adams, 1992; Lemoine et al., 1995).

Under saturating red light, photosynthesis of E. siliculosus follows a circadian rhythm, with maxima at about noon, and can be stimulated by a pulse of blue light (Schmid & Dring, 1992). This stimulation is also observed in other Phaeophytes, but mostly absent in green or red algae (Schmid et al., 1994b). Blue light induces several responses: an acidification at the surface of E. siliculosus (Schmid & Dring, 1993), accompanied by bicarbonate uptake (Schmid, 1998). A plasma membrane H+-ATPase is thought to be activated, and the resulting acidification to increase the conversion of inline image to CO2 in the extracellular space (Schmid & Dring, 1993). Recently, such blue light-induced H+-ATPases have been identified in the brown alga Laminaria digitata (Klenell et al., 2002). Blue light also triggers the mobilization of an internal carbon source, since photosynthesis is stimulated even in the absence of external Ci (Schmid & Dring, 1996). As a result, a C4-like metabolism was initially proposed to exist in E. siliculosus (Schmid & Dring, 1996) but not all the enzymes necessary for a C4 cycle were detected (Busch & Schmid, 2001) and the pool of intermediates seems to be too small to act as an organic carbon stock (Hillrichs & Schmid, 2001). The sequestration of a pool of Ci in the vacuole and its movement to the cytosol in response to blue light is now the favoured hypothesis (Schmid & Hillrichs, 2001). Analysis of the sequence of the E. siliculosus genome will help to confirm whether or not a C4 pathway exists in the brown algae.

In contrast to the Plantae (Moreira et al., 2000), the Phaeophyceae do not store the carbon assimilated by photosynthesis as insoluble starch granules, but instead as the soluble 1,3-β-glucan polymer (laminarin) localized in the cytosol (Craigie, 1974), and as mannitol, involved in osmo-acclimation (Davis et al., 2003; see Section V.2: Abiotic stresses). Brown algae also produce complex polysaccharides which constitute their cell wall. They synthesize some neutral polysaccharides in common with land plants, such as cellulose (Carpita & McCann, 2000), but also unique anionic polysaccharides, such as alginates and sulphated fucans (Kloareg & Quatrano, 1988). Ectocarpus has not been well studied in this respect, but preliminary analyses in our laboratory confirm that all of the polysaccharides typical of brown algae are present in this genus (Estelle Deniaud, pers. comm.). The biosynthetic pathways of these brown algal polysaccharides are essentially unknown and the genome of E. siliculosus will be a much-anticipated asset to investigate these crucial metabolisms.

2. Lipid metabolism

Worldwide, at least 50 species of brown algae are used as human food. Their lipid content has therefore attracted considerable attention from the viewpoint of both nutrition and pharmacology. A number of studies have been conducted to profile and quantify the fatty acids and the different classes of lipids in these organisms, and to investigate whether the various lipid patterns correlate with the taxonomic position or any other characteristic of the brown algae. Brown algal polar lipids include several common glycolipids (monogalactosyldiacylglycerol, digalastosyldiacylglycerol, sulfoquinovosyldiacylglycerol) and phospholipids (phosphatidylcholine, phosphatidylethanolamine, phosphatidylinositol, phospahtidylglycerol and diphosphatidylglycerol). Interestingly, several reports have highlighted the high proportion of long-chain polyunsaturated fatty acids (LC-PUFAs) (Eichenberger et al., 1993). Moreover, most of the Phaeophyceae contain the betaine lipids diacylglycerylhydroxymethyl-N,N,N-trimethyl-β-alanine (DGTA) and diacylglyceryltrimethylhomoserine, and either contain the common phosphatidylcholine phospholipids in surprisingly low amounts or do not produce them at all (Eichenberger et al., 1993). All Ectocarpales contain phosphatidylcholine, but exhibit interspecific variation in their DGTA content: E. fasciculatus strains contain this lipid while E. siliculosus strains do not (Müller, 1995; Müller & Eichenberger, 1995). Diacylglycerylhydroxymethyl-N,N,N-trimethyl-β-alanine may therefore be used as a taxonomic marker (Müller, 1995), although an E. fasciculatus strain deficient in DGTA biosynthesis has been described. Genetic analysis of such a strain identified an autosomal locus necessary for the biosynthesis of this lipid (Müller & Eichenberger, 1997).

A novel phosphoglyceride, designated PX, was first isolated from E. siliculosus. It was shown to account for 2–4% of total lipids (Schmid et al., 1994c), and to accumulate mostly in the plasma membrane of gametes. Since PX is rich in 20:4n-6 (arachidonic acid) and 20:5n-3 (eicosapentaenoic acid), it has been suggested that it could represent a potential reservoir for pheromone precursors. PX was subsequently detected in other brown algae (Ectocarpales, Fucales and Sphacelariales; Schmid et al., 1994c).

Lipid metabolism plays also major roles in the control of defense mechanisms. Therefore, together with the genome sequence, it is likely that the efforts to screen mutants with altered resistance to pathogen attacks will lead to the phenotyping of plants impaired in lipid or fatty acid metabolic pathways, thereby potentially revealing novel specific traits of this metabolism in brown algae.

V. Interactions with the environment

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

1. Ectocarpus pathogens

Despite their small size and ephemeral life stages, filamentous brown algae have been frequently reported to be plagued by various pathogens, including viruses (Müller et al., 1998) and eukaryotic parasites of different phylogenetic lineages: oomycetes, chytrids and hyphochytrids (Andrews, 1976; Küpper & Müller, 1999; Müller et al., 1999) and by parasites related to the Plasmodiophorea (Karling, 1944; Maier et al., 2000). In addition, numerous historical records described ectocarpoids with abnormal sporangia or vegetative cells suspected to contain unknown parasites (Rattray, 1885; Müller et al., 1998).

The oomycete Eurychasma dicksonii has been described mainly in wild populations of Pylaiella littoralis (Küpper & Müller, 1999), but it displays a broad host range and infects various brown algae, including Ectocarpus (Müller et al., 1999), in which it was initially described by Wright (1879). There is a current effort to set up a defined pathosystem using E. siliculosus and E. dicksonii, and Ectocarpus strains have been shown to exhibit differential susceptibility to a defined Eurychasma strain. Conversely, several Eurychasma strains exhibit different host specificities, suggesting coevolution of the two species (Gachon et al., 2007). The molecular bases of resistance and virulence are under investigation.

Chytrids were described earlier by Petersen (1905), and the hyphochytrid Anisolpidium ectocarpii was described by Karling (1943) and Johnson (1957). Like E. dicksonii, Chytridium polysiphoniae (Chytridiomycota) is ubiquitous and can infect many hosts, including E. siliculosus and E. fasciculatus (Müller et al., 1999). Interestingly, its negative effects on photosynthesis of its host was described at the cellular level in the related ectocarpoid P. littoralis using fluorescence kinetic microscopy (Gachon et al., 2006). Recently, the 18S rRNA genes of Chytridium polysiphoniae and Eurychasma dicksonii were sequenced and used to clarify their phylogenetic affiliations (Küpper et al., 2006). The plasmodiophorean Maullinia ectocarpii is an obligate intracellular parasite of Ectocarpus spp. (Maier et al., 2000). However, the extent to which this infection occurs in nature and its effect on algal fitness are presently unknown.

Viral infections represent by far the most studied phenomenon in E. siliculosus (Müller, 1996; Müller & Knippers, 2001). Until the late 1980s, most reports of virus infections in brown algal tissues were based on electron microscopy studies, which sporadically described ‘virus-like particules’ (VLPs). Viruses were obtained in culture for the first time from a New Zealand strain of E. siliculosus after lysis of host cells, allowing evaluation of their infection potential (Müller, 1991; Müller et al., 1990). Virus infections were found in approx. 50% of the individuals of a given natural population (Dixon et al., 2000; Müller et al., 2000) and were shown to occur worldwide in correlation with the cosmopolitan distribution of E. siliculosus (Müller, 1991; Sengco et al., 1996).

The viruses that infect different ectocarpoid algae exhibit considerable variability in size and diameter and, in general, display a high degree of host specificity (Müller et al., 1998). However, several instances of trans-specific infection have been described, for example between EsV-1 (Ectocarpus siliculosus virus-1) and Kuckuckia kylinii (Müller, 1992; Müller & Schmid, 1996) and also between EfasV-1 (Ectocarpus fasciculatus virus-1) and E. siliculosus (Müller et al., 1996; Sengco et al., 1996). Interestingly, EsV-1 and EfasV-1 are the most similar of the brown algal viruses in terms of their genome size (Müller et al., 1996).

The EsV-1 virus specifically infects the single-celled gametes or spores, that is, the only cells in the life history that lack a cell wall (Maier & Müller, 1998). Following infection, a single copy of the viral DNA appears to integrate into the host genome (Delaroque et al., 1999). The viral DNA is then transmitted, via mitotic divisions, to all the cells of the developing alga. This has been confirmed by regenerating algae from protoplasts derived from virus-infected gametophytes (Kuhlenkamp & Müller, 1994). Despite the fact that they carry the integrated virus, vegetative cells do not produce viral particles (Müller et al., 1998). Viral particles are only produced in reproductive organs (sporangia and gametangia) of mature algae from where they are released to infect a new generation of zoids. In addition to these cycles of re-infection, the viral genome can also be transmitted to progeny through meiosis, in which case it segregates as a Mendelian factor and is inherited by half of the progeny (Müller, 1991; Bräutigam et al., 1995). The pathogenic character of viral infections has been unambiguously confirmed, but this association's main impact is on reproductive success. Plant sterility varies from partial (Müller et al., 1990) to total (Müller & Frenzer, 1993), but no significant difference in photosynthesis, respiration and growth rate were observed in infected gametophytes or sporophytes (Del Campo et al., 1997). This contrasts with the reduced photosynthetic performance of Feldmannia species infected with FsV (Robledo et al., 1994).

The EsV-1 genome is a circular DNA molecule of a relatively large size (335 kbp) for a phycodnavirus (Van Etten & Meints, 1999; Van Etten et al., 2002) with double-stranded regions interrupted by single-stranded regions (Lanka et al., 1993; Klein et al., 1994). Both EsV-1 and the related Feldmania irregularis virus (FirrV-1) have been sequenced (Delaroque et al., 2001, 2003). EsV-1 contains approx. 231 genes with a wide range of predicted functions, including DNA metabolism, signalling, transposition, DNA integration and polysaccharide metabolism (Delaroque et al., 2000a,b, 2003). It has also been proposed that the ability of the virus to integrate into its host's genome could be exploited to develop a transformation vector for a wide range of brown algae, including E. siliculosus (Henry & Meints, 1994, Delaroque et al., 1999). However, the complex integration pattern of the virus into the algal genome will considerably complicate this task (N. Delaroque, pers. comm.). A microarray has been constructed to analyse EsV-1 gene expression (Declan Schroeder, pers. comm.) and it will be particularly interesting in the future to couple the analysis of viral and genome-wide host gene expression during viral infection.

The development of genomic tools provides a new context to investigate the possible genetic basis of the coevolution between some pathogens and brown algae. The search for inducers of defense responses and resistance against parasites is also still ongoing as, in contrast to kelps, Ectocarpus does not react with an oxidative burst upon recognition of alginate fragments (Küpper et al., 2002a).

2. Abiotic stresses

Ectocarpus siliculosus is able to exploit a wide range of habitats and environmental conditions (see Section II.2: Distribution). This feature seems likely to be based at least as much as on a high intrinsic genetic variability as on a general physiological toughness, as illustrated by work carried out on copper and saline stress responses.

Interspecific variations in copper tolerance have been observed between different strains of E. fasciculatus and E. siliculosus, with the latter being the most tolerant (Morris, 1974). Differences have also been observed among E. siliculosus strains that are differently exposed to copper in their natural habitat (Russell & Morris, 1970; Hall, 1981). Cu2+ interferes with the general process of photosynthesis in brown algae, and particularly in E. siliculosus, by competing with magnesium for metal-binding sites in the chlorophyll molecules (Küpper et al., 2002b). A study of the mechanism of tolerance to copper and other heavy metals suggested a co-tolerance to copper, cobalt and zinc, and provided evidence for an exclusion mechanism to explain the particularly low sensitivity of E. siliculosus copper-tolerant strains (Hall et al., 1979; Hall, 1980, 1981). However, as yet there is no clear explanation for the intraspecific variation with respect to this trait within this species.

It has been suggested that the ability of some E. siliculosus strains to tolerate copper may be useful for the development of bioassays in which this alga is used for monitoring marine antifouling characteristics of copper-based materials (Hall & Baker, 1985, 1986). Copper chloride has been used to inhibit E. siliculosus infestations in tank cultures of Gracilaria gracilis (Van Heerden et al., 1997).

Russell & Bolton (1975) reported the occurrence of salinity ecotypes within E. siliculosus. This study was extended by Thomas & Kirst (1991a,b), who showed that large differences in photosynthesis, accumulation of osmotically active compounds (mannitol; Davis et al., 2003) and vitality occur between E. siliculosus isolates from different geographic locations following changes in salinity. They also observed that sporophytes were more salt-tolerant than gametophytes, irrespective of their level of ploidy.

Detailed investigations are necessary to decipher the physiological and cellular bases of salt and heavy metal tolerance in E. siliculosus. Mutagenesis and transcriptomic approaches will thus help to better understand the mechanisms involved in osmotic and oxidative adaptation, and to explain how these algae can cope with such a wide range of environmental conditions.

VI. Conclusion

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

Taken together, the above sections illustrate the broad range of phenomena that have been studied in Ectocarpus and provide an indication of the domains that could be further explored in the future. Notably, a large proportion of this past work, covering many diverse aspects of Ectocarpus biology, was carried out in Dieter Müller's laboratory in Konstanz, and the efforts of this group have therefore laid the foundations for the development of Ectocarpus as a model organism.

The Ectocarpus genome project has federated a consortium of laboratories with an interest in this organism and these laboratories are currently developing several molecular tools. These include mutant screens, genetic transformation and genome-scale analysis of gene expression. Several developmental mutants have been isolated and positional cloning of some of the affected genes should be feasible in the near future. The availability of the genome sequence together with the ability to analyse gene function by forward and reverse genetic approaches will make it possible to address additional questions, many of which have been evoked in this review. Examples include the biosynthesis of diverse, brown algal-specific metabolites, such as lipids and complex cell wall components, and the genetic basis of resistance to biotic and abiotic aggression. The genome will also be an invaluable aide for the study of the ecology of Ectocarpus, by serving as the base for the development of neutral and selected molecular markers for the analysis of field isolates. In conclusion, the Ectocarpus genome sequence and the tool development associated with this project are providing access to a relatively unexplored branch of the eukaryotic tree and some exciting discoveries can be expected in this domain in the coming years.


  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References

We are grateful to N. Delaroque (Max Planck Institute in Jena) for authorization to communicate unpublished information about viral integration, and to Declan Schroeder (Marine Biological Association in Plymouth) for authorization to mention the microarray for the EsV-1 virus.


  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Diversity and taxonomy, distribution and ecology
  5. III. Development
  6. IV. Metabolism
  7. V. Interactions with the environment
  8. VI. Conclusion
  9. Acknowledgements
  10. References
  • Alberte RS, Friedman AL, Gustafson DL, Rudnick MS, Lyman H. 1981. Light-harvesting systems of brown algae and diatoms. Isolation and characterization of chlorophyll a/c and chlorophyll a/fucoxanthin pigment-protein complexes. Biochimica et Biophysica Acta 635: 304316.
  • Andrews JA. 1976. The pathology of marine algae. Biological Reviews 51: 211253.
  • Baker JRJ, Evans LV. 1973a. The ship fouling alga Ectocarpus. I. Ultrastructure and cytochemistry of plurilocular reproductive stages. Protoplasma 77: 113.
  • Baker JRJ, Evans LV. 1973b. The ship fouling alga Ectocarpus. II. Ultrastructure of unilocular reproductive stages. Protoplasma 77: 181189.
  • Baldauf SL. 2003. The deep roots of eukaryotes. Science 300: 17031706.
  • Barrett J, Anderson JM. 1980. The P-700-chlorophyl alpha-protein complex and two major light-harvesting complexes of Acrocarpia paniculata and other brown seaweeds. Biochimica et Biophysica Acta 590: 309323.
  • Beer S. 1994. Mechanisms of inorganic carbon acquisition in marine macroalgae. Progress in Phycological Research 10: 179207.
  • Berthold G. 1881. Die geschlechtliche Fortpflanzung der eigentlichen Phaeosporeen. Mitteilungen aus der Zoologischen Station zu Neapel 2: 401413.
  • Boalch GT. 1961. Studies on Ectocarpus in culture. Journal of the Marine Biological Association. 41: 279304.
  • Boland W, König WA, Krebber R, Müller DG. 1989. Separation of enantiomeric algal pheromones and related hydrocarbons by gas-liquid chromatography on modified cyclodextrins as chiral stationary phases. Biosynthetic relevance of racemic by-products. Helvetica Chimica Acta 72: 12881292.
  • Boland W, Marner FJ, Jaenicke L. 1983. Comparative receptor study in gamete chemotaxis of the seaweeds Ectocarpus siliculosus and Cutleria multifida – An approach to interspecific communication of algal gametes. European Journal of Biochemistry 134: 97103.
  • Boland W, Pohnert G, Maier I. 1995. Biosynthesis of algae pheromones. 4. Pericyclic-reaction in nature – spontaneous cope rearrangement inactivates algae pheromones. Angewandte Chemie 34: 16021604.
  • Bolton JJ. 1983. Ecoclinal variation in Ectocarpus siliculosus (Phaeophyceae) with respect to temperature growth optima and survival limits. Marine Biology 73: 131138.
  • Bouck GB. 1965. Fine structure and organelle associations in brown algae. Journal of Cell Biology 26: 523537.
  • Bräutigam M, Klein M, Knippers R, Müller DG. 1995. Inheritance and meiotic elimination of a virus genome in the host Ectocarpus siliculosus (Phaeophyceae). Journal of Phycology 31: 823827.
  • Busch S, Schmid R. 2001. Enzymes associated with β-carboxylation in Ectocarpus siliculosus (Phaeophyceae): are they involved in net carbon acquisition? European Journal of Phycology 36, 6170.
  • Cardinal A. 1964. Étude sur les Ectocarpacées de la Manche. Nova Hedwigia 15: 186.
  • Carpita N, McCann M. 2000. The cell wall. In: BuchananB, GruissemW, JonesR, eds. Biochemistry and molecular biology of plants. American Society of Plant Physiologists. Rockville, MD, USA: 52108.
  • Cho, GY, Lee SH, Boo SM. 2004. A new brown algal order, Ishigeales (Phaeophyceae), established on the basis of plastid protein-coding rbcL, psaA, and psbA region comparisons. Journal of Phycology 40: 921936.
  • Corellou F, Coelho SM, Bouget FY, Brownlee C. 2005. Spatial re-organisation of cortical microtubules in vivo during polarisation and asymmetric division of Fucus zygotes. Journal of Cell Science 118 :27232734.
  • Craigie JS. 1974. Storage products. In: StewartWPD, ed. Algal physiology and biochemistry. Berkeley, CA, USA: University of California Press, 206235.
  • Davis TA, Volesky B, Mucci A. 2003. A review of the biochemistry of heavy metal biosorption by brown algae. Water research 37: 43114330.
  • Davis, RH. 2004. The age of model organisms. Nature Reviews Genetics 5: 6975.
  • De Martino A, Douady D, Quinet-Szely M, Rousseau B, Crépineau F, Apt K, Caron L. 2000. The light-harvesting antenna of brown algae: highly homologous proteins encoded by a multigene family. European Journal of Biochemistry 267: 55405549.
  • Del Campo E, Ramazanov Z, Garcia-Reina G, Müller DG. 1997. Photosynthetic responses and growth performance of virus-infected and noninfected Ectocarpus siliculosus (Phaeophyceae). Phycologia 36: 186189.
  • Delaroque N, Boland W, Müller DG, Knippers R. 2003. Comparisons of two large phaeoviral genomes and evolutionary implications. Journal of Molecular Evolution 57: 613622.
  • Delaroque N, Maier I, Knippers R, Müller DG. 1999. Persistent virus integration into the genome of its algal host, Ectocarpus siliculosus (Phaeophyceae). Journal of General Virology 80: 13671370.
  • Delaroque N, Müller DG, Bothe G, Pohl T, Knippers R, Boland W. 2001. The complete DNA sequence of the Ectocarpus siliculosus Virus EsV-1 genome. Virology 287: 112132.
  • Delaroque N, Wolf S, Müller DG, Knippers R. 2000a. The brown algal virus EsV-1 particle contains a putative hybrid histidine kinase. Virology 273: 383390.
  • Delaroque N, Wolf S, Müller DG, Knippers R. 2000b. Characterization and immunolocalization of major structural proteins in the brown algal virus EsV-1. Virology 269: 148155.
  • Demmig-Adams B, Adams III WW. 1992. Photoprotection and other responses to high light stress. Annual Review of Plant Physiology and Plant Molecular Biology 43: 599626.
  • Dillwyn LW. 1809. British Confervae; or colored figures and descriptions of the British plants referred by botanists to the genus Conferva. London, UK: W. Phillips, 187, 1–6 (Index and Errata), Plates 69, 100–109, A–G (with text).
  • Dixon NM, Leadbeater BSC, Wood KR. 2000. Frequency of viral infection in a field population of Ectocarpus fasciculatus (Ectocarpales, Phaeophyceae). Phycologia 39: 258263.
  • Draisma SGA, Peters AF, Fletcher RL. 2003. Evolution and taxonomy in the Phaeophyceae: effects of the molecular age on brown algal systematics. In: NortonTA, ed. Out of the past. Collected reviews to celebrate the jubilee of the British Phycological Society. Belfast, UK: The British Phycological Society, 87102.
  • Eichenberger W, Araki S, Müller DG. 1993. Betaine lipid and phospholipids in brown algae. Phytochemistry 34: 13231333.
  • Evans LV. 1966. Distribution of pyrenoids among some brown algae. Journal of Cell Science 1: 449454.
  • Gachon CM, Day JG, Campbell CN, Proschold T, Saxon RJ, Küpper FC. in press. Culture collection of algae and protozoa (CCAP): a biological resource for protistan genomics. Gene.
  • Gachon CM, Müller DG, Gaj G, Küpper FC. 2007. Confronting disease: not all algae are equal. The Phycologist 75: 15 (abstract).
  • Gachon CMM, Küpper H, Küpper FC, Šetlík I. 2006. Witnessing effects of a pathogen on photosynthesis of its host at the cellular level: chlorophyll fluorescence kinetic microscopy of Pylaiella littoralis (Phaeophyceae) infected by Chytridium polysiphoniae (Chytridiomycota). European Journal of Phycology 41: 395403.
  • Geissler U. 1983. Die salzbelastete Fluβstrecke der Werra-ein Binnenlandstandort für Ectocarpus confervoides (Roth) Kjellman. Nova Hedwigia 37: 193217.
  • Geller A, Müller DG. 1981. Analysis of the flagellar beat pattern of male Ectocarpus siliculosus gametes (Phaeophyta) in relation to chemotactic stimulation by female cells. Journal of Experimental Biology 92: 53666.
  • Grossman AR, Bhaya D, Apt KE, Kehoe DM. 1995. Light-harvesting complexes in oxygenic photosynthesis: diversity, control, and evolution. Annual Review of Genetics 29: 231288.
  • Hall A. 1980. Heavy-metal co-tolerance in a copper-tolerant population of the marine fouling alga, Ectocarpus siliculosus (DILLW) LYNGBYE. New Phytologist 85: 7378.
  • Hall A. 1981. Copper accumulation in copper-tolerant and non-tolerant populations of the marine fouling alga Ectocarpus siliculosus (Dillw.) Lyngbye. Botanica Marina 24: 223228.
  • Hall A, Baker AJM. 1985. Settlement and growth of copper-tolerant Ectocarpus siliculosus (DILLW.) LYNGBYE on different copper-based antifouling surfaces under laboratory conditions.1. Corrosion trials in seawater and development of an algal culture system. Journal of Materials Science 20: 11111118.
  • Hall A, Baker AJM. 1986. Settlement and growth of copper-tolerant Ectocarpus siliculosus (DILLW.) LYNGBYE on different copper-based antifouling surfaces under laboratory conditions. 2. A comparison of the early stages of fouling using light and electron microscopy. Journal of Materials Science 21: 12401252.
  • Hall A, Fielding AH, Butler M. 1979. Mechanism of copper tolerance in the marine fouling alga Ectocarpus siliculosus– evidence for an exclusion mechanism. Marine Biology 54: 195199.
  • Hartmann M. 1934. Untersuchungen über die Sexualität von Ectocarpus siliculosus. Archiv für Protistenkunde 83: 110163.
  • Henry EC, Meints RH. 1994. Recombinant viruses as transformation vectors of marine macroalgae. Journal of Applied Phycology 6: 247253.
  • Hillrichs S, Schmid R. 2001. Activation by blue light of inorganic carbon acquisition for photosynthesis in Ectocarpus siliculosus: organic acid pools and short-term carbon fixation. European Journal of Phycology 36: 7179.
  • Johnson TW. 1957. Resting spore development in the marine phycomycete Anisolpidium ectocarpii. American Journal of Botany 44: 875878.
  • Karling JS. 1943. The life history of Anisolpidium ectocarpii gen. nov. et sp. nov., and a synopsis and classification of other fungi with anteriorly uniflagellate zoospores. American Journal of Botany 30: 637648.
  • Karling JS. 1944. Phagomyxa algarum n. gen.n. sp., an unusual parasite with plasmodiophoralean and proeomyxean characteristics. American Journal of Botany 31: 3852.
  • Katsaros C, Kreimer G, Melkonian M. 1991. Localization of tubulin and a centrin-homologue in vegetative cells and developing gametangia of Ectocarpus siliculosus (Dillw.) Lyngb. (Phaeophyceae, Ectocarpales). Botanica Acta 104: 8792.
  • Kawai H. 1988. A flavin-like autofluorescent substance in the posterior flagellum of golden and brown algae. Journal of Phycology 24: 114117.
  • Kawai H. 1992. A summary of the morphology of chloroplast and flagellated cells in the Phaeophyceae. Korean Journal of Phycology 7: 3343.
  • Kawai H, Hanyuda T, Draisma SGA, Müller DG. 2007. Molecular phylogeny of Discosporangium mesarthrocarpum (Phaeophyceae) with a reinstatement of the order Discosporangiales. Journal of Phycology 43: 186194.
  • Kawai H, Müller DG, Fölster E, Häder DP. 1990. Phototactic responses in the gametes of the brown alga, Ectocarpus siliculosus. Planta 182: 292297.
  • Kim HS, Lee IK. 1992. Morphotaxonomic studies on the Korean Ectocarpaceae (Phaeotphyta) I. Genus Ectocarpus Lyngbye. Korean Journal of Phycology 7(2): 225242.
  • Klarzynski O, Plesse B, Joubert J-M, Yvin J-C, Kopp M, Kloareg B, Fritig B. 2000. Linear β-1,3 glucans are elicitors of defense responses in tobacco. Plant Physiology 124: 10271037.
  • Klein M, Lanka STJ, Müller DG, Knippers R. 1994. Single-stranded regions in the genome of the Ectocarpus siliculosus virus. Virology 202: 10761078.
  • Klenell M, Snoeijs P, Pedersén M. 2002. The involvement of a plasma membrane H+-ATPase in the blue-light enhancement of photosynthesis in Laminaria digitata (Phaeophyta). Journal of Phycology 38: 11431149.
  • Kloareg B, Quatrano RS. 1988. Structure of the cell walls of marine algae and ecophysiological functions of the matrix polysaccharides. Oceanography and Marine Biology. An Annual Review. 26: 259315.
  • Knight M. 1929. Studies in the Ectocarpaceae. II. The life-history and cytology of Ectocarpus siliculosus, Dillw. Transactions of the Royal Society of Edinburgh 56: 307332.
  • Kornmann P. 1956. Über die Entwicklung einer Ectocarpus confervoides-Form. Pubblicazioni Della Stazione Zoologica di Napoli 28: 3243.
  • Kornmann P, Sahling PH. 1977. Meeresalgen von Helgoland. Benthische Grün-, Braun- und Rotalgen. Helgoländer Wissenschaftliche Meeresuntersuchungen 29: 1289.
  • Kreimer G, Kawai H, Müller DG, Melkonian M. 1991. Reflective properties of the stigma in male gametes of Ectocarpus siliculosus (Phaeophyceae) studied by confocal laser scanning microscopy. Journal of Phycology 27: 268276.
  • Kuhlenkamp R, Müller DG. 1994. Isolation and regeneration of protoplasts from healthy and virus-infected gametophytes of Ectocarpus siliculosus (Phaeophyceae). Botanica Marina 37: 525530.
  • Küpper FC, Maier I, Müller DG, Loiseaux-de Goër S, Guillou L. 2006. Phylogenetic affinities of two eukaryotic pathogens of marine macroalgae, Eurychasma dicksonii and Chytridium polysiphoniae. Cryptogamie Algologie 27: 165184.
  • Küpper FC, Müller DG. 1999. Massive occurrence of the heterokont and fungal parasites Anisolpidium, Eurychasma and Chytridium in Pylaiella littoralis (Ectocarpales, Phaeophyceae). Nova Hedwigia 69: 381389.
  • Küpper FC, Müller D, Peters A, Kloareg B, Potin P. 2002a. Oligoalginate recognition and oxidative burst play a key role in natural and induced resistance of sporophytes of Laminariales. Journal of Chemical Ecology 28: 20572081.
  • Küpper H, Šetlík I, Spiller M, Küpper FC, Prášil O. 2002b. Heavy metal-induced inhibition of photosynthesis: targets of in vivo heavy metal chlorophyll formation. Journal of Phycology 38: 429441.
  • Lanka STJ, Klein M, Ramsperger U, Müller DG, Knippers R. 1993. Genome structure of a virus infecting the marine brown alga Ectocarpus siliculosus. Virology 193: 802811.
  • Le Gall Y, Brown S, Marie D, Mejjad M, Kloareg B. 1993. Quantification of nuclear DNA and G-C content in marine macroalgae by flow cytometry of isolated nuclei. Protoplasma 173: 123132.
  • Lemoine Y, Harker M, Rmiki NE, Rousseau B, Berkaloff C, Duval JC, Young AJ, Britton G. 1995. Xanthophyll cycle operation and photoprotection in brown algae: effects of high light and desiccation. In: MathisP, ed. Photosynthesis, from light to biosphere, vol. IV. Dordrecht, the Netherlands: Kluwer, 119122.
  • Lofthouse PF, Capon B. 1975. Ultrastructural changes accompanying mitosporogenesis in Ectocarpus parvus. Protoplasma 84: 8399.
  • Lyngbye HC. 1819. Tentamen hydrophytologiae danicae continens omnia hydrophyta cryptogamma Daniae, Holsatiae, Faeroae, Islandiae, Groendlandiae hucusque cognita, systematice disposita, descripta et iconibus illustrata, adjectis simul speciebus norvegicis. Opus, praemio. Copenhagen, Denmark: Schultz.
  • Maier I. 1997a. The fine structure of the male gamete of Ectocarpus siliculosus (Ectocarpales, Phaeophyceae). I. General structure of the cell. European Journal of Phycology 32: 241253.
  • Maier I. 1997b. The fine structure of the male gamete of Ectocarpus siliculosus (Ectocarpales, Phaeophyceae). II. The flagellar apparatus. European Journal of Phycology 32: 255266.
  • Maier I, Müller DG. 1998. Virus binding to brown algal spores and gametes visualized by DAPI fluorescence microscopy. Phycologia 37: 6063.
  • Maier I, Parodi E, Westermeier R, Müller DG. 2000. Maullina ectocarpii gen. et sp. nov. (Plasmodiophorea), an intracellular parasite in Ectocarpus siliculosus (Ectocarpales, Phaeophyceae) and other filamentous brown algae. Protist 151: 225238.
  • Maier I, Schmid CE. 1995. An immunofluorescence study on lectin binding sites in gametes of Ectocarpus siliculosus (Ectocarpales, Phaeophyceae). Phycological Research 43: 3342.
  • McHugh DJ. 2003. A guide to the seaweed industry. FAO Fisheries Technical Paper no. 441. Rome: FAO, 105 pp.
  • Moreira D, Le Guyader H, Philippe H. 2000. The origin of red algae and the evolution of chloroplasts. Nature 405: 6972.
  • Morris OP. 1974. Inter-specific differences in responses to copper by natural populations of Ectocarpus. British Phycology Journal 9: 269272.
  • Morris OP, Russel G. 1974. Inter-specific differences in responses to copper by natural populations of Ectocarpus. British Phycology Journal 9: 269272.
  • Müller DG. 1963. Die Temperaturabhängigkeit der Sporangienbildung bei Ectocarpus siliculosus von verschiedenen Standorten. Pubblicazioni Della Stazione Zoologica di Napoli 33: 310314.
  • Müller DG. 1964. Life-cycle of Ectocarpus siliculosus from Naples, Italy. Nature 26: 1402.
  • Müller DG. 1966. Untersuchungen zur Entwicklungsgeschichte der Braunalge Ectocarpus siliculosus aus Neapel. Planta 68: 5768.
  • Müller DG. 1967. Generationswechsel, Kernphasenwechsel und Sexualität der Braunalge Ectocarpus siliculosus im Kulturversuch. Planta 75: 3954.
  • Müller DG. 1972a. Life cycle of the brown alga Ectocarpus fasciculatus var. refractus (Kütz.) Ardis. (Phaeophyta, Ectocarpales) in culture. Phycologia 11: 1113.
  • Müller DG. 1972b. Studies on reproduction in Ectocarpus siliculosus. Société Botanique Française Mémoires 1972: 8798.
  • Müller DG. 1976a. Sexual isolation between a European and an American population of Ectocarpus siliculosus (Phaeophyta). Journal of Phycology 12: 252254.
  • Müller DG. 1976b. Quantitative evaluation of sexual chemotaxis in two marine brown algae. Zeitschrift für Planzenphysiologie 80: 120130.
  • Müller DG. 1978. Locomotive responses of male gametes to the species specific sex attractant in Ectocarpus siliculosus (Phaeophyta). Archiv für Protistenkunde 120: 371377.
  • Müller DG. 1980. Entwicklung von Ectocarpus siliculosus (Phaeophyta). Film C 1308 des IWF, Göttingen 1979. Publikationen zu wissenschaftlichen Filmen, Sektion Biologie, Serie 13 11/C 1308: 115.
  • Müller DG. 1991. Mendelian segregation of a virus genome during host meiosis in the marine brown alga Ectocarpus siliculosus. Journal of Plant Physiology 137: 739743.
  • Müller DG. 1992. Intergeneric transmission of a marine plant DNA virus. Naturwissenschaften 79: 3739.
  • Müller DG. 1995. Taxonomic value of a betaine lipid in Ectocarpus (Phaeophyceae) and the first record of a sexual population of E. fasciculatus for the Pacific Ocean. Phycological Research 43: 175177.
  • Müller DG. 1996. Host-virus interactions in marine brown algae. Hydrobiologia 327: 2128.
  • Müller DG, Eichenberger W. 1995. Crossing experiments, lipid composition, and the species concept in Ectocarpus siliculosus and E. fasciculatus (Phaeophyceae, Ectocarpales). Journal of Phycology 31: 173176.
  • Müller DG, Eichenberger W. 1997. Mendelian genetics in brown algae: inheritance of a lipid defect mutation and sex alleles in Ectocarpus siliculosus (Ectocarpales, Phaeophyceae). Phycologia 36: 7981.
  • Müller DG, Falk H. 1973. Flagellar structure of the gametes of Ectocarpus siliculosus (Phaeophyta) as revealed by negative staining. Archiv für Mikrobiologie 91: 313322.
  • Müller DG, Frenzer K. 1993. Virus infection in three marine brown algae: Feldmannia irregularis, F. simplex, and Ectocarpus siliculosus. Hydrobiologia 260–261: 3744.
  • Müller DG, Jaenicke L, Donike M, Akintobi T. 1971. Sex attractant in a brown alga: chemical structure. Science 171: 815817.
  • Müller DG, Kapp M, Knippers R. 1998. Viruses in marine brown algae. Advances in Virus Research 50: 5067.
  • Müller DG, Kawai H, Stache B, Lanka STJ. 1990. A virus infection in the marine brown alga Ectocarpus siliculosus (Phaeophyceae). Botanica Acta 103: 7282.
  • Müller DG, Knippers R. 2001. Phaeovirus, phycodnaviridae. In: TidonaCA, DaraiG, eds. The springer index of viruses. Berlin, Germany: Springer, 732736.
  • Müller DG, Küpper FC, Küpper H. 1999. Infection experiments reveal broad host ranges of Eurychasma dicksonii (Oomycota) and Chytridium polysiphoniae (Chytridiomycota), two eukaryotic parasites in marine brown algae (Phaeophyceae). Phycological Research 47: 217223.
  • Müller DG, Maier I, Müller H. 1987. Flagellum autofluorescence and photoaccumulation in heterokont algae. Photochemistry and Photobiology 46: 10031008.
  • Müller DG, Schmid CE. 1988. Qualitative and quantitative determination of pheromone secretion in female gametes of Ectocarpus siliculosus (Phaeophyceae). Biological Chemistry Hoppe-Seyler 369: 647653.
  • Müller DG, Schmid CE. 1996. Intergeneric infection and persistence of Ectocarpus virus DNA in Kuckuckia (Phaeophyceae, Ectocarpales). Botanica Marina 39: 401405.
  • Müller DG, Sengco M, Bräutigam M, Schmid CE, Kapp M, Knippers R. 1996. Comparison of two DNA viruses infecting the marine brown algae Ectocarpus siliculosus and E. fasciculatus. Journal of General Virology 77: 23292333.
  • Müller DG, Westermeier R, Morales J, Garcia Reina G, Del Campo E, Correa JA, Rometsch E. 2000. Massive prevalence of viral DNA in Ectocarpus (Phaeophyceae, Ectocarpales) from two habitats in the North Atlantic and South Pacific. Botanica Marina 43: 157159.
  • Oliveira L, Bisalputra T. 1973. Studies in the brown alga Ectocarpus in culture. Journal of Submicrobial Cytology 5: 107120.
  • Oliveira L, Bisalputra T. 1977a. Ultrastructural studies in the brown alga Ectocarpus in culture: autolysis. New Phytologist 78: 139145.
  • Oliveira L, Bisalputra T. 1977b. Ultrastructural studies in the brown alga Ectocarpus in culture: ageing. New Phytologist 78: 131138.
  • Oltmanns F. 1899. Ueber die Sexualität der Ectocarpeen. Flora 86: 8699.
  • Papenfuss GF. 1935. Alternation of generations in Ectocarpus siliculosus. Botanical Gazette 96: 421446.
  • Pedersén M. 1968. Ectocarpus fasciculatus: marine brown alga requiring kinetin. Nature 218: 776.
  • Pedersén M. 1973. Identification of a cytokinin, 6-(3 methyl-2-butenylamino) purine, in sea water and the effect of cytokinins on brown algae. Physiologia Plantarum 28: 101105.
  • Pedersén PM. 1989. Studies on Kuckuckia spinosa (Fucophyceae, Sorocarpaceae): life history, temperature gradient experiments, and synonymy. Nordic Journal of Botany 9: 443447.
  • Peters AF, Marie D, Scornet D, Kloareg B, Cock JM. 2004a. Proposal of Ectocarpus siliculosus (Ectocarpales, Phaeophyceae) as a model organism for brown algal genetics and genomics. Journal of Phycology 50: 10791088.
  • Peters AF, Scornet D, Müller DG, Kloareg B, Cock JM. 2004b. Inheritance of organelles in artificial hybrids of the isogamous multicellular chromist alga Ectocarpus siliculosus (Phaeophyceae). European Journal of Phycology 39: 235242.
  • Petersen HE. 1905. Contributions à la connaissance des phycomycètes marins (Chytridinae Fischer). Oversigt over det kgl. Danske Videnskabernes Selskats Forhandlinger 5: 439488.
  • Rattray J. 1885. Note on Ectocarpus. Transactions of the Royal Society of Edinburgh 32: 589602.
  • Ravanko O. 1970. Morphological, developmental, and taxonomic studies in the Ectocarpus complex (Phaeophyceae). Nova Hedwigia 20: 79252.
  • Robledo DR, Sosa PA, Garcia-Reina G, Müller DG. 1994. Photosynthetic performance of healthy and virus-infected Feldmannia irregularis and F. simplex (Phaeophyceae). European Journal of Phycology 29: 247251.
  • Rousseau F, De Reviers B. 1999. Circumscription of the order Ectocarpales (Phaeophyceae): bibliographical synthesis and molecular evidence. Cryptogamie, Algologie 20: 518.
  • Russell G. 1966. The genus Ectocarpus in Britain. I. The attached forms. Journal of the Marine Biological Association of the United Kingdom 46: 267294.
  • Russell G. 1967a. The genus Ectocarpus in Britain. II. The free-living forms. Journal of the marine biological Association of the United Kingdom 47: 233250.
  • Russell G. 1967b. The ecology of some free-living Ectocarpaceae. Helgoländer wissenschaftliche Meeresuntersuchungen 15: 155162.
  • Russell G. 1983a. Parallel growth patterns in algal epiphytes and Laminaria blades. Marine Ecology Progress Series 13: 303304.
  • Russell G. 1983b. Formation of an ectocarpoid epiflora on blades of Laminaria digitata. Marine Ecology Progress Series 11: 181187.
  • Russell G, Bolton JJ. 1975. Euryhaline ecotypes of Ectocarpus siliculosus (Dillw.) Lyngb. Estuarine and Coastal Marine Science 3: 9194.
  • Russell G, Morris OP. 1970. Copper tolerance in the marine fouling alga Ectocarpus siliculosus. Nature 228: 288289.
  • Sauvageau C. 1896. Sur la conjugaison des zoospores de l’Ectocarpus siliculosus. Comptes rendus de l’Académie des Sciences 123: 431433.
  • Sauvageau C. 1897. La copulation isogamique de l’Ectocarpus siliculosus est-elle apparente ou réelle? Mémoires de la Société Nationale des Sciences Naturelles de Cherbourg 30: 293304.
  • Schmid CE. 1993. Cell-cell-recognition during fertilization in Ectocarpus siliculosus (Phaeophyceae). Hydrobiologia 260–261: 437443.
  • Schmid CE, Müller DG, Eichenberger W. 1994c. Isolation and characterization of a new phospholipid from brown algae. Intracellular localization and site of biosynthesis. Journal of Plant Physiology 143: 570574.
  • Schmid CE, Schroer N, Kawai H, Müller DG. 1992. Isolation and biochemical characterization of different gamete membranes in the chromophyte alga Ectocarpus siliculosus (Phaeophyceae). Plant Physiology and Biochemistry 30: 703712.
  • Schmid CE, Schroer N, Müller DG. 1994a. Female gamete membrane glycoproteins potentially involved in gamete recognition in Ectocarpus siliculosus (Phaeophyceae). Plant Science 102: 6167.
  • Schmid R. 1998. Photosynthesis of Ectocarpus siliculosus in red light and after pulses of blue light at high pH – evidence for bicarbonate uptake. Plant, Cell & Environment 21: 523529.
  • Schmid R, Dring MJ. 1992. Circadian rhythm and fast responses to blue light of photosynthesis in Ectocarpus (Phaeophyta, Ectocarpales). Planta 187: 5359.
  • Schmid R, Dring MJ. 1993. Rapid, blue-light-induced acidifications at the surface of Ectocarpus and other marine macroalgae. Plant Physiology 101: 907913.
  • Schmid R, Dring MJ. 1996. Influence of carbon supply on the circadian rhythmicity of photosynthesis and its stimulation by blue light in Ectocarpus siliculosus: clues to the mechanism of inorganic carbon acquisition in lower brown algae. Plant, Cell & Environment 19: 373382.
  • Schmid R, Dring MJ, Forster RM. 1994b. Kinetics of blue light stimulation and circadian rhythmicity of light-saturated photosynthesis in brown algae: a species comparison. Journal of Phycology 30: 612621.
  • Schmid R, Hillrichs S. 2001. Uptake and accumulation of inorganic carbon in Ectocarpus siliculosus and its relation to blue light stimulation of photosynthesis. European Journal of Phycology 36: 257264.
  • Sengco M, Bräutigam M, Kapp M, Müller DG. 1996. Detection of virus DNA in Ectocarpus siliculosus and E. fasciculatus (Phaeophyceae) from various geographic areas. European Journal of Phycology 31: 7378.
  • Silva PC, Basson P, Moe RL. 1996. Catalogue of the benthic marine algae of the Indian Ocean. University of California Publications in Botany 79: 11259.
  • Stache, B. 1990. Sexual compatibility and species concept in Ectocarpus siliculosus (Ectocarpales, Phaeophyceae) from Italy, North Carolina, Chile, and New Zealand. In: Garbary, DJ, SouthRG eds. Evolutionary biogeography of the marine algae of the North Atlantic. Berlin, Germany: Springer Verlag, 173186.
  • Stache-Crain B, Müller DG, Goff LJ. 1997. Molecular systematics of Ectocarpus and Kuckuckia (Ectocarpales, Phaeophyceae) inferred from phylogenetic analysis of nuclear and plastid-encoded DNA sequences. Journal of Phycology 33: 152168.
  • Thomas DN, Kirst GO. 1991a. Salt tolerance of Ectocarpus siliculosus (Dillw.) Lyngb.: comparison of gametophytes, sporophytes and isolates of different geographic origin. Botanica Acta 104: 2636.
  • Thomas DN, Kirst GO. 1991b. Differences in osmoacclimation between sporophytes and gametophytes of the brown alga Ectocarpus siliculosus. Physiologia Plantarum 83: 281289.
  • Van Etten JL, Graves MV, Müller DG, Boland W, Delaroque N. 2002. Phycodnaviridae – large DNA algal viruses. Archives of Virology 147: 14791516.
  • Van Etten JL, Meints RH. 1999. Giant viruses infecting algae. Annual Review of Microbiology 53: 447494.
  • Van Heerden PDR, Robertson BL, De Kock L. 1997. Inhibition of Ectocarpus siliculosus infestations with copper chloride in tank cultures of Gracilaria gracilis. Journal of Applied Phycology 9: 255259.
  • West JA, Kraft GT. 1996. Ectocarpus siliculosus (Dillwyn) Lyngb. from Hopkins River Falls, Victoria – the first record of a freshwater brown alga in Australia. Muelleria 9: 2933.
  • Wiencke C, Clayton MN. 2002. Antarctic seaweeds. Synopses of the Antarctic benthos, Vol. 9. Ruggell, Liechtenstein: Gantner.
  • Wright EP. 1879. On a species of Rhizophydium parasitic on species of Ectocarpus, with notes on the fructification of the Ectocarpi. Transactions of the Royal Irish Academy 26: 369379.