What we really know about the composition and function of microalgae cell coverings? - an overview

ABSTRACT

Cell coverings can be observed in all major groups of organisms, which include animals, plants, fungi, protists and prokaryotes. They play a key role in assuring cell survival or adaptation to certain environmental conditions. Since the term algae refers to a polyphyletic and very artificial group, the cell coverings of these organisms are very diverse in molecular composition and with different arrangements. Differences have taxonomic value since they allow microalgae phyla or even minor taxonomic groups, such as classes, orders or families, to be distinguished. Understanding the structure of cell coverings is also fundamental for the use of microalgae to obtain products of commercial value. Despite its importance, the composition and architecture of microalgae coverings is still poorly understood, especially considering the great diversity of organisms. Diatom frustules are the most studied coverings due their uses in areas of bio- and nanotechnology. There is a lack of information about the cell wall, lorica, periplast, amphiesma and scales. This study is a review with the aim of synthesizing literature information on microalgae cell coverings to describe their compositions, arrangements, functions and industrial uses.

Keywords:
algal coverings variety; biological interactions; cell surface; molecular structures; taxonomical value

Introduction

Microalgae comprise eukaryotic and procariotic organisms (cyanobacteria) (Gigova & Marinova 2016), commonly studied together due to their similar photoautotrophic metabolism (Saad & Atia 2014). They show many differences in cell structure and physiology due to their polyphyletic origin and also because they evolved to adapt to many different environments, such as freshwater, seawater, salt lakes, soil, arctic environments, deserts (Raja et al. 2014; Zancan et al. 2006) and even in association with other organisms such as corals, plants and fungi (in lichens) (Sanders 2001; Wooldridge 2013). Among the particularities used to characterize the different groups of microalgae and cyanobacteria there are the cell coverings, which are the special structures that surround their cells.

The many types of cell coverings are related to the needs of different microalgae, which include interactions with chemical substances, connection between cells, fixation in substrates, protection, communication, reproduction and maintenance of the cell shape (Peterson & Quie 1981; Hoson 2002; Okuda 2002; Yoshimi et al. 2017). Microalgae cell coverings have been studied for decades, but details about their ultrastructure and composition are not well known, especially when compared to the higher availability of information about other structures, or even about pigments and reserve substances. Due to economic interests, the studies are focused on diatoms, chlorophytes and charophytes, with some generalizations being established for these phyla. There is a lack of recent studies about the cell coverings of other phyla. Further, generalizations have low value even for the well-known phyla since studies are concentrated in few species although species-specific particularities are commonly reported (Domozych et al. 2012).

Here, we present a review to synthesize the available information about composition, architecture and function of the different types of cell coverings present in cyanobacteria and microalgae.

Types and composition of cyanobacteria and microalgae coverings

The cell coverings of autotrophic microorganisms have different names according to their particular structure, position in relation to cell surface and chemical composition (Okuda 2002). Table 1 synthesizes information about these coverings types, their major compounds and taxonomical groups in which they are present.

Table 1
Types and composition of cell covering found in algae and cyanobacteria.

Cell covering	Class	Major compounds	References
Cell wall	Cyanophyceae	Peptidoglycans	Rudolf et al. 2015
Amphiesma	Dinophyceae	Cellulose	Chan et al. 2019;
Periplast	Cryptophyceae	Proteins	Hoef-Emden & Melkonian 2003; Brett et al. 1994
Pellicle	Euglenophyceae	Proteinaceous strips	Cavalier-Smith 2017; Sommer 1965
Lorica		Mucopolysaccharides and minerals	Poniewozik 2017
Coccolith (calcified scale)	Hapthophyceae	Calcium carbonate	Walker et al. 2018; Faber & Preisig 1994
Frustule	Bacillariophyceae	Silica	Tommasi et al. 2017; Nakajima & Volcani 1969
Lorica	Chrysophyceae	Chitin and cellulose	Herth et al. 1977; Herth & Zugenmaier 1979
Silica scales	Chrysophyceae	Silica	Leadbeater & Barker 1995
	Sinurophyceae	Silica	Sandgren & Hall 1996
Cell wall	Chlorophyceae	Cellulose	Okuda 2002
Organic scales		2-keto sugar acids	Becker et al. 1994
Cell Wall	Charophyceae	Cellulose	Okuda 2002

Some coverings are located internally in relation to the plasma membrane. They are found in Dinophyta, Cryptophyta and Euglenophyta and are called amphiesma, periplast and pellicle, respectively (Gantt 1971; Morrill & Loeblich 1983; Leander et al. 2001). Cyanobacteria and some microalgae have coverings that are located externally the plasma membrane. Even the cell walls of Cyanobacteria and the eukaryotic green algae (Chlorophyta and Charophyta) being different in structure and composition, they are an example of external coverings with similar function. Other external structures are the lorica present in some Euglenophyta and Ochrophyta, the scales of some Ochrophyta and Haptophyta and the frustules of Bacillariophyta, which is one of the most particular algal coverings. Details on each covering type are presented in the following items.

Inner cell coverings

Amphiesma

Dinoflagellates can be divided into two major groups: naked organisms with no thick coverings and armored organisms, which have an amphiesma (Gómez 2007). This term (from Greek, amphi = around, esthma = clothing) was coined by Schütt (1895) and refers to the complete covering of armored dinoflagellates, which includes the plasma membrane as the outermost layer (Sekida et al. 2004; Morrill 1984), a layer of membranous vesicles, which may contain glucan tecal plates, and a pellicle (Pozdnyakov & Skarlato 2012) (Fig. 1). The term theca can be also used (Dodge & Crawford 1970), but not as a synonym for amphiesma since it refers only to the layer formed by the vesicles containing thecal plates in armored (thecate) dinophytes. The thecal plates are described originally as a cellulosic structure (Swift & Remsen 1970; Okuda & Sekida 2007), but a recent study (Wang et al. 2011) showed that proteins are also present. The number and disposition of thecal plates are important features for dinoflagellates classification (Dodge 1983). These membranous vesicles are empty or contain amorphous materials in athecate dinoflagellates (Dodge & Crawford 1970).

Figure 1
Schematic representation of a cell of Dinophyta (A), showing its two typical flagella. Representation of the structural components of the amphiesma (B): The pellicle layer; (C) Outer plate membrane; (D) Techal vesicle; (E) Techal plate; (F) Cytoplasmatic membrane and the outermost membrane (G). Adapted from Wang et al. (2011).

The innermost layer is a membrane that some authors consider as a part of the amphiesma, but others (see Morrill & Loeblich 1983) consider it as a pellicle. This layer is present in some species and contains sporopollenin-like substances that confers resistance to it (Morrill & Loeblich 1983; Okuda 2002). In some athecate dinoflagellates, the pellicle may be the most important layer to confer resistance to the cell surface, maintaining the cell’s shape (Saldarriaga & Taylor 2017).

The amphiesma is a dynamic structure that undergoes many changes throughout the life cycle of the organisms (Sekida et al. 2004; Pozdnyakov & Skarlato 2012). Despite all published studies, the structure and genesis of the amphiesma remain not fully understood (Pozdnyakov & Skarlato 2012). Sekida et al. (2001) showed that the vesicles are formed in the non-motile phase of the life cycle and after that the thecal plates are formed inside them in the motile phase.

Periplast

The cryptophytes have an asymmetric cell shape with clearly defined dorsi-ventral/right-left sides (Hoef-Emden & Melkonian 2003) that has taxonomic significance. This shape partially results of the presence of a vestibule, which is a subapical invagination of plasma membrane, but it is mainly related to a rigid periplast, which is the typical cell covering in this phylum (Brett et al. 1994). It covers the entire cell, except the flagella and the vestibular/gullet region (Perasso et al. 1997). The vestibule (from where the flagella emerge) can extend internally to form a gullet or continue along the ventral surface to form a furrow (Kugrens & Lee 1991).

The periplast of cryptophytes is composed of two proteic layers, the inner periplast component (IPC) and the surface periplast component (SPC), with the plasma membrane sandwiched between them (Gantt 1971; Brett et al. 1994) (Fig. 2). Nevertheless, there are also some species that have a simpler periplast composed by only the plasma membrane and the inner layer (Kugrens & Lee 1987). The totality of characteristics and functions of the periplast are uncertain, but stiffness, flexibility and elasticity are commonly attributed to it (Faust 1974). A fourth possible function is to protect the integrity of the cell membrane during the explosive discharges of the ejectisomes, a type of extrusive organelles (Hausmann 1978; 1979).

Figure 2
A-Scheme of a cell of Cryptophyta. B-Representation of structural components of the periplast: Inner periplast component (IPC), surface periplast component (SPC) and the plasma membrane (PM). Adapted from Brec et al. (1999).

The morphology and organization of the periplast are different among Cryptophyceae and more than one type of IPC were described (Brett & Wetherbee 1986). The IPC develops within specific regions called anamorphic zones that are located around the vestibule (Brett & Wheterbee 1996a). The IPC is able to grow throughout the life cycle, allowing the elongation and expansion of the cell (Brett & Wheterbee 1996a). Depending of the taxon, the IPC is formed by a unique continuous layer or it could be formed by several scales arranged internally in relation to the plasma membrane (Brett et al. 1994). The SPC may appear as dense mats of an unidentified fibrillar material, complex rosulate scales or highly ordered surface plates (Brett & Wheterbee 1986; Brett et al. 1994). The microarchitecture of these plates were described in detail by Brett & Wheterbee (1996b), who showed that these plates are formed by aligned tiny subunits. Studies suggest that these subunits of SPC are produced in the Golgi apparatus and secreted through the endomembrane system to be added at the edges of the periplast (Brett et al. 1994; Perasso et al. 1997).

The periplast is a complex and unique type of cell covering and some researchers dedicated their work to elucidate its formation, structure and composition by using refined techniques such as immunocytochemistry (Perasso et al. 1997), scanning electron microscopy, freeze-fracture and freeze-etch (See review: Brett et al. 1994). However, even with the necessity to better understand the periplast structure and functioning, there are few recent studies focused on this type of cell covering. As can be seen in this review, relevant researches are dated from the 1970s to the 1990s.

Pellicle

Although the pellicle can be found among the dinophytes, it is much more complex in Euglenophyta. It is the most important covering in this phylum and is the most rigid structure in the cell surface of most species. The pellicle of euglenophytes can be described as a complex region containing proteinaceous strips, microtubules and tubular cisternae of endoplasmic reticulum that runs along the length of the cell beneath the plasma membrane (Leedale 1964; Sommer 1965; Schwelitz et al. 1970; Vismara et al. 2000; Strother et al. 2019) (Fig. 3).

Figure 3
Scheme of a cell of Euglenophyta (A). Magnification of the pellicle (B) showing the strips (C), the pores (D) and the spaces between the strips (E). Schematic representation of the pellicle’s components and organization (F-J). The plasma membrane (F) covers the strips (G) that are connected by the centrines (H). In J and I the endoplasmic reticulum and the microtubules are represented respectively. Adaptaded from Cavalier Smith (2017) and Leander et al. (2001).

The strips are considered the major component of the euglenids pellicles, and their general ultrastructure is relatively well understood (Strother et al. 2019). They are composed by proteins named articulins which are arranged in parallel and result in a typical ultrastructure that can be used to differentiate species or genera (Cavalier-Smith 2017). The quantity of strips has taxonomic value since it varies widely among species, but is conserved within them (Cavalier-Smith 2016). Another important aspect of strips is how they are organized in cell surface, since they can be arranged in longitudinal rows or helically twisted (Leander et al. 2007). When arranged longitudinally, the strips make the pellicle rigid and prevent changes in the cell's shape, as observed in the most primitive euglenoids that form the classes Entosiphonea, which has fewer strips (12 or less), Stavomonadea and Ploeotarea (superclass Rigimonada) (Cavalier-Smith 2016; Cavalier-Smith 2017). The pellicles with helical strips are found in euglenoids of the superclass Spirocuta, comprising heterotrophic Peranema Stein, 1859, and ancestrally photosynthetic Euglenophyceae, which in turn have several strips (14-80) (Cavalier-Smith 2016; Cavalier-Smith 2017). These pellicles show higher malleability and are often associated with a peculiar mode of cellular locomotion called metabolic or "euglenoid movement" (Leander et al. 2001).

At the junctions between the strips there are the pellicle pores. They are small openings whose function is to provide access for two different ejectile organelles (muciferous bodies and mucocysts) to the cell surface (Leander et al. 2001). The strips are connected each other by oblique traversing fibres of centrin, which is a calcium-sensitive contractile protein closely related to the body's reorientations during photophobic responses and flagellar contractions (Höhfeld et al. 1988). Intimately associated with each strip there is still the cisterna of endoplasmic reticulum, that pump and store calcium for release when centrin contractions are required (Cavalier-Smith 2017).

Extracellular coverings

Cyanobacterial cell wall

Cyanobacteria are a special type of bacteria since they are the only group of prokaryotes that can perform oxygenic photosynthesis (Zhang et al. 2018). They are ubiquitous organisms mainly due to their adaptation to various types of environments and their tolerance to extreme conditions (Gaysina et al. 2019). Their cell wall is part of their adaptive success.

Bacteria are generally classified as gram positive or negative according to the chemical and physical properties of their cell walls (Hiremath & Bannigidad 2011). Cyanobacteria are gram-negative bacteria, with the cell wall located externally to the plasma membrane. This wall consists of a peptidoglycan layer that is involved by a superficial layer, also called outer membrane (Fig. 4). The plasma membrane is also commonly called inner membrane by some authors, who consider it as a third layer composing the cell wall (Silhavy et al. 2010). The outer membrane is a particularity of Gram-negative bacteria and it is formed by an asymmetric bilayer, in which the inner face is composed of phospholipids, while the outer face is composed of lipopolysaccharides (LPS) (Zhang et al. 2013) that play a key role in bacterial pathogenicity (Maldonado et al. 2016). The outer membrane is a selective permeation barrier (Nikaido 2003) that is involved in cell nutrition and also confers resistance to a variety of detergents and antibiotics (Doerrler 2006).

Figure 4
A- Representation of a filamentous Cyanobacteria. B-Scheme showing the components of the gram-negative cell wall of Cyanobacteria. In this membrane are located some integral proteins (C) and porins (D). The outermost layer is composed of lipopolysaccharides (LPS) (E) that are found on the surface of the outermost plasma membrane (F). Just below there is a peptidoglycan layer (G), composed of pentaglycine cross-links (a) and alternating polymers of N-acetylmuramic acid and N-acetylglucosamine (b). Below this layer is another layer of phospholipid membrane (H). Adapted from Aiad et al. (2016).

The peptidoglycan layer is composed by repeated units of the disaccharide N-acetyl glucosamine and by N-acetyl muramic acid, which are cross-linked by pentapeptide side chains (Vollmer et al. 2008). This layer gives rigidity to the cyanobacterial cells, maintaining their shape. It also confers a protection against differences in osmotic pressure between the external and internal media, and also serves as a scaffold for anchoring proteins and teichoic acids (See review: Irazoki et al. 2019). Despite its rigidity, the peptide glycan layer is sufficiently dynamic to allow cell growth, division and morphogenesis (Zhang et al. 2018).

Although the general structure of cyanobacterial cell walls is the same observed in gram-negative bacteria, some characteristics of gram-positive walls and other particularities are also present (Hoiczyk & Hansel 2000). Their peptidoglycan layer, for example, is considerably thicker (reaching 700 nm in larger species, like Oscillatoria princeps Gomont 1892) than those observed in most gram-negative bacteria (5-10 nm) (Hoiczyk & Hansel 2000). In Synechocystis Sauvageau, 1892, the degree of cross-linking between peptidoglycan chains is greater than that usually found in heterotrophic gram-negative bacteria and is more similar to the reported values for gram-positive bacteria (Hoiczyk & Hansel 2000). Further, cyanobacteria cell walls have components that are absent in the cell walls of other gram-negative bacteria. For example, they have carotenoids (Resch & Gibson 1983) and the fatty acid b-hydroxypalmitic as a substitute for the hydroxymyristic acid commonly found in other gram-negative bacteria (Jurgens & Weckesser 1985).

Eukariotic microalgae cell wall

Among the microalgae coverings, the term “cell wall” is reserved for a thick, rigid and continuous structure that is mainly composed of cellulose microfibrils (Okuda 2002). This kind of cell covering is found in the green algae (Chlorophyta and Charophyta). The molecular structure of the cellulose of these algae is the same of the cellulose found in plants, which is a linear polymer of β-(1-4)-linked d-glucose (Baldan et al. 2001; Fry 2003). However, the structure of the cellulosic walls is not the same for the both groups (Okuda 2002), except for few algal taxa more related to plants (Sørensen et al. 2011; Domozych et al. 2012). Beyond cellulose, other polymers like xylans and mannans may also occur as the major structural polymer in some algae (Okuda 2002) and some species of Chlorophyta have no cellulose (Imam 1985).

The structural polymers are embedded in an amorphous matrix composed by polysaccharides, which varies among different green algae taxa (Domozych et al. 2012). Then, together with cellulose, hemicellulose, pectins, and other polysaccharides composing this matrix were described in cell walls of various microalgae (Sørensen et al. 2011, see Tab. 2) (Fig. 5). However, there is a lack of information about the complete structure and composition of algae cell walls. Recently, more attention has been paid to algal cell wall due to the new tools and techniques allowing to do very detailed studies and mainly due to the need to better understand these structures. This knowledge is fundamental to solve questions about the morphology and physiology of these organisms and their interaction with the environment. It is also necessary for the development of methods to disrupt cells for the extraction of various compounds of economic interest (Baudelet et al. 2017), such as pigments and fatty acids.

Figure 5
A- Representation of a colony of Chlorophyta. B- Magnification showing the main components of the green algae cell wall. In C is represented the layer of pectic compounds in which the microfibrils of cellulose (D) and hemicellulose (E) are immersed.

Table 2
Chemical composition of green-algae cell walls.

Phylum	Cell wall componentes	References
Charophyta	Uronic acids	Popper & Fry 2003
	Cellulose, hemicelluloses and pectins	Domozych et al. 2010; Sørensen et al. 2011
	Pectins	Domozych et al. 2007; Eder & Lütz-Meindl 2008; Eder & Lütz-Meindl 2010; Domozych et al. 2011
	Cellulose	Wurdack 1923; Herburger & Holzinger 2015
	AGPs and hemicelluloses	Eder et al. 2008
	Chitin	Wurdack 1923
Clorophyta	Cellulose	Parker 1964; Yamada & Sakaguchi 1982; Baldan et al. 2001; Němcová 2003
	Glycoproteins	Goodenough & Heuser 1985
	Sporopollenin	He et al. 2016
	Aminoacids and saccharides	Abo-Shady et al. 1993
	HRGP and hemicelluloses	Estevez et al. 2009
	Cellulose and other polysacharides	Piro et al. 2000
	Chitin	Wurdack 1923
	D-glucose, D-mannose	Parker 1964
		Wurdack 1923
	Pectins	Bisalputra & Weier 1963; Yamada & Sakaguchi 1982

Organic scales

Members of Prasinophyceae (Chlorophyta) have their cells covered by organic scales. These scales are mainly composed of acid polysaccharides (2-keto sugars), with some proteins being present in lesser amount (Becker et al. 1994). Prasinophyceae scales are synthesized in the Golgi cisterns and transported to the cell surface by exocytosis (Moestrup & Walne 1979). Interestingly, a cell can show many types of scales arranged in several layers (1-5) on the surface of the cell body and even of the flagella (Becker et al. 1994) (Fig. 6). Less common, it is also observed the scales fused in one piece, as occurs in the genera Tetraselmis Stein, 1878, and Scherffelia Pascher, 1912 (Arora et al. 2013). In the order Pyramimonadales the scales are arranged in 3 layers. These are considered the most complex among the Prasinophyceae. In the innermost layer, the scales are small, square or pentagonal; in the middle layer the scales are naviculoid, have the form of a spider web or the form of a box, while the scales of the outer layer have the shape of a crown (Daugbjerg 2000). The scales morphometry is widely varied and very important as a taxonomic character to differentiate between orders, families and genera (Becker et al. 1994).

Figure 6
Some types of organic scales found in Pyramimonas diskoicola. Adapted from Harðardóttir et al. 2014.

Some haptophytes (such as Phaeocystis Scherffel, 1899, Prymnesium Carter, 1937, Pavlova Butcher, 1952, and Chrysochromulina Lackey, 1939) have their cells covered with organic scales (Young & Henriksen 2003). Composed by cellulose, these scales are produced in the Golgi apparatus and transported through vesicles to the cell surface (Jordan & Chamberlain 1997). In some cases, they can also cover the haptonema or one of the flagella (Vargas et al. 2007). Scales morphology varies among taxa within this phyllum and then these structures are commonly used as a taxonomic character (Eikrem et al. 2017). In Pavlovophyceae, for example, the scales are structurally simpler and have a knoblike form, while in the Prymnesiophyceae, the scales are more ornamented and shaped like plates (Vargas et al. 2007). For many species, the organic scales serve as a calcification matrix for the formation of rigid scales that are named coccoliths (Houdan et al. 2004; Liu et al. 2010). These structures are special scales that are only observed in Haptophyta and a detailed description will be presented in the next item.

Coccoliths

Coccoliths are the most common cell covering found in haptophytes. They are calcified plates (CaCO₃ as calcite) that cover the cells forming a coccosphere (Taylor et al. 2016; Müller 2019) (Fig. 7). The arrangement of these plates is a taxonomic character that is used for even distinguish organisms in the species level (Chrétiennot-Dinet et al. 2014). The coccoliths are formed in the cisterns of the dicytiosomes (Manton 1966) and are released to the cell surface by fusion of the plasmalema with the cisternal membrane (Eikrem et al. 2017).

Figure 7
Schematic representation of a coccolithophore (A) and an amplification of the structural unit that form its covering, the coccolith (B).

There are two main types of coccoliths, the heterococcoliths and the holococcoliths, based on their ultrastructure and morphology (Braarud et al. 1995). Some possible functions attributed to coccoliths are protection against predation and virus attack (Monteiro et al. 2016), optimization of light absorption by the cell (Young 1994), dissipation of excessive absorbed light energy to avoid photo damage under nutrient limitation (Paasche 2002), regulation of buoyancy (Young 1994) and carbon concentration mechanism (Sikes et al. 1980). However, these hypotheses have not yet been proven (Eikrem et al. 2017) and none of them has sufficient and consistent evidence to be scientifically accepted (Müller 2019).

The species of coccholite-coated haptophyta are commonly called coccolithophore. The oldest recorded coccolithophores are from the upper Triassic sediments, approximately 225 Ma. (Bown et al. 2004). They were and are abundant in the marine phytoplankton and show a historical and current very important role in carbon cycling. Biomineralization of coccolithophores controls the alkalinity, chemistry of photic zone carbonates of the oceans, and the carbonate precipitation (through the calcification reaction) is a short-term source of CO₂ to the high ocean and atmosphere (Vargas et al. 2007).

Lorica

The particularity of lorica in relation to other coverings is that it is not adhered to the plasma membrane, being similar to an envelope or armor. Lorica can be found in some Euglenophyta and Ochrophyta. For Euglenophyta, pellicle is their typical covering but some genera such as Strombomonas Deflandre, 1930, Trachelomonas Ehrenberg, 1834, and Ascoglena Stein, 1978, additionally have lorica (Duangjan & Wolowski 2013). This covering is a rigid and mucilaginous protective envelope composed by mucopolysaccharides and minerals (mainly iron and sometimes manganese) (Poniewozik 2017) that surround the cell and have a gap from which the flagellum emerges (Fig. 8A). The lorica surface can be smooth, but it usually presents granular or rough appearance due to the agglutination of particles from the environment. The lorica shape and its ornamentation are very important taxonomic characters to differentiate genera and species among the euglenophytes (Brosnan et al. 2005). Lorica can be colorless, but they generally have a yellow-brown or orange color due to the impregnation of minerals (Leedale 1975).

Figure 8
Schematic representation of some types of Euglenophyta lorics (A). A Dinobryon colony and a detailed vision of the main regions (B): The cup (C), the stalk (D) and the foot (E). Adapted from Conforti (2010).

Although chrysophyceans (Ochrophyta) lorica are similar to euglenophytes lorica in relation to minerals impregnation, colors and microarchitecture (Dunlap et al. 1987), their coverings are mainly composed of chitin and cellulose (Herth & Zugenmaier 1979). In some Chrysophyceae, the organization of lorica can be simplified in foot, stalk and cup (Fig. 8B). These structures have species-specific features (see Peck 2010) with evident taxonomic significance, such as shape, size and ornamentation (Belcher 1969; Kapustin 2019). Composition and architecture are also important (Dunlap et al. 1987). Dinobryon Ehrenberg, 1834, for example, has a lorica with a vase or beaker-shaped form, while Chrysococcus Klebs, 1892, has globular and Lagynium present a flask-shaped lorica (Kristiansen & Škaloud 2017). As observed in euglenophytes, manganese and iron compounds can be present and are responsible for the dark and opaque color of some chrysophycean lorica (Dunlap et al. 1987).

The formation of a new lorica is not well understood for many species, but it is better understood for Dinobryon. It was observed that the formation of a new lorica in this genus begins after cell division. The daughter cell moves to the edge near the opening of the parent lorica, where it connects and fixes. After that, it will first secret the small basal cone and then the complete cup-shape of the lorica (Karim & Round 1967).

Silica scales

Chrysophyceans (Ochrophyta) of the family Paraphysomonadaceae, mainly the genera Chrysosphaerella Lauterborn, 1896, and Paraphysomonas De Saedeleer, 1930, do not have lorica, but silicified scales covering their cells (Kristiansen 2008). They are attached outside to the plasma membrane (Němcová & Pichrtová 2012) with no defined pattern. Silica scales are radially or biradially symmetrical and their sizes vary from about 1 to 10 μm (Škaloud et al. 2013).

The scales of chrysophyceans have an endogenous origin and are formed inside a vesicle of deposition of silica, which is in turn derived from the endoplasmic reticulum (Lee 2008). The scales are extruded from the cell and placed in the correct position on its surface (Kristiansen 1986). The covering formed by scales is a dynamic structure that allows the addition of new scales during both growth and division (Škaloud et al. 2013). Techniques of electron microscopy allowed to know much about the structure of the scales, which is highly variable among species (Kristiansen & Škaloud 2017) and therefore have taxonomic significance. However, a basic structure is common for all species, and it can be described as a perforated plate that can have ribs, spines and other ornaments (Kristiansen 1986).

Silica scales are also found in Synurophyceae (Ochrophyta) cells. They are formed internally in silica deposition vesicles and then they are transported to the cell surface (Wee 1997). Interestingly, several scale types can occur on the same cell and each type show a particular distribution on the cell surface (Neustupa et al 2010; Škaloud et al. 2012) (Fig. 9). The genus Synura Ehrenberg, 1834, for example, has three distinct scales by cell: body, apical and rear scales that are characterized by their different length to width ratios (Škaloud et al. 2012). Abiotic factors in the environment like pH (Siver 1989) and temperature (Řezáčová-Škaloudová et al. 2010) seems to have some influence on the morphological differentiation of the scales. The morphology is specie-specific and have highlighted taxonomy significance (Kristiansen 2002), especially the body scales, that exhibit the most highly developed and complex characters (Škaloud et al. 2012).

Figure 9
Scheme of silica scales found in the genus Mallomonas. Scale found in the species M. flora (A), M. matvienkoae (B), M. ouradion ( C). Adapted from Peck (2010).

Frustule

Diatoms are one of the most easily recognizable groups among the algae due to the presence of its characteristic silicified cell wall covering (Reimann et al. 1965). This special cell wall is named frustule and is formed by two parts of similar size that are called valves. One valve is slightly larger than the other, with the smaller valve fitting inside the larger one. This fitting is connected by structures called girdle bands that allow preciselly link these valves around the protoplasm (Kröger & Poulsen 2008; Tesson & Hildebrand 2010). The large valve is named epitheca and the smaller is the hypotheca (Fig. 10). A symmetrical structure of the leaflets divides the diatoms into two generic groups: centric, with radial symmetry and as pennates with bilateral symmetry, sometimes with a transverse groove, the raphe.

Figure 10
A-Scheme of a closed frustule of a penate diatom. B- The main components of the frustule: The epitheca (C), the girdle bandes (D) and the hypotheca (E).

The frustule is mainly composed by silica, that gives rigidity to its structure, but it is in association with an organic wall composed by proteins, polyamines and polysaccharides (Nakajima &Volcani 1969; Kröger et al. 1999; Gügi et al. 2015). The organic matter seems to play a role in cell adhesion to surfaces and protection to cell desiccation (Kröger & Poulsen 2008). Four families of proteins have been described in diatom’s cell walls: frustulins, pleuralins, p150 family and silaffins (Kröger & Poulsen 2008). Silaffins are suggested to be the molecule involved in silica formation (Kröger & Poulsen 2008). In relation to the carbohydrates (mono or polysaccharides), there are many types that were observed in frustule (Tab. 3). Chitin, for example, is a polysaccharide that was detected in association with silica (Gügi et al. 2015). A detailed study of Phaeodactylum tricornutum Bohlin, 1898, showed the presence of glucuronic acid and mannose (Ford & Percival 1965). These carbohydrates are common in high amounts in frustules of several species of diatoms while the quantity of fucose and xylose is more variable (Gügi et al. 2015).

Table 3
Main components of the frustule.

Chemical component			References
Inorganic	silica		Nakajima &Volcani 1969
Organic	Proteins	Frustulins	Kröger et al. 1994; 1996; 1997; 1999; 2000; 2001; Kröger & Poulsen 2008
		p150 family
		Silaffins
		Pleuralins
	Polyamines		Kröger et al. 1999
	Monosaccharides	Rhamnose	Hecky et al. 1973
		Fucose	Hecky et al. 1973 Gügi et al. 2015
		Ribose	Hecky et al. 1973
		Arabinose	Hecky et al. 1973
		Xylose	Hecky et al. 1973 Gügi et al. 2015
		Mannose	Hecky et al. 1973 Ford & Percival 1965
		Galactose	Hecky et al. 1973
		Glucose	Hecky et al. 1973
		Glucuronic acid	Ford & Percival 1965
	Polysaccharide	Chitin	Gügi et al. 2015

The knowledge on frustule composition, structure and synthesis has not only taxonomical importance, but this cell covering has also commercial value and many industrial uses. The ease of cultivation in artificial environments (culture media) and the availability of fossilized frustules (diatomite) make diatom silica a promising natural alternative to synthetic materials for biomedical, environmental, agricultural, and energy applications (Maher et al. 2018; Terraciano et al. 2018). Diatoms have been currently studied for biotechnological and nanotechnological purposes, being involved in techniques of nanofabrication, chemo and biosensor, classification and control of particles in micro and Nano fluid (Jamali et al. 2012). Dolatabadi & La-Guardia (2011) for example, present in their review the applications of silicious diatoms and nanomaterials in biosensing (drug and gene delivery) and their utility to form complex metallic nanostructures.

Coverings affect commercial exploration of microalgae: limitations and tools to disrupt cells and assess products

Algal cell walls are one of the main products of exploitation among marine macroscopic algae, from which sulfated polysaccharides and other compounds are extracted and used in a wide variety of industrial segments (Jönsson et al. 2020). In relation to the microscopic ones, diatom frustules, as already mentioned, have been studied for use in biotechnological and nanotechnological purposes (Jamali et al. 2012). Other products not related to covering composition are also of commercial interest. Algae, and especially microalgae, has been increasingly targeted as a sustainable source of high added value compounds used by the industry of pharmaceuticals, cosmetics and nutrition, and some are also alternative feedstocks for biofuel production (Wu et al. 2017; Dixon & Wilken 2018). Various microalgae and cyanobacteria are known to produce these targeted compounds, but commercial exploration and research are concentrated in few genera, as summarized in the Table 4.

Table 4
Most commonly cultivated microalgae and cyanobacteria for commercial purposes or for researches to support a possible future exploration. The major compounds of interest by genus or species are also presented.

Phyllum	Genus or species	Compounds of interest	References
Chlorophyta	Botryococcus	Hydrocarbons	Metzger & Largeau 2005
	Chlorella	Pigments and lipids	Safi et al. 2014
	Scenedesmus		Wiltshire et al. 2000
	Dunaliella	β-Carotene	Raja et al. 2007
	Haematococcus	Astaxanthin	Shah et al. 2016
	Tetraselmis	Eicosapentaenoic acid (EPA), Vitamin E	Pereira et al. 2019
Haptophyta	Isochrysis galbana	Fatty acids	Bandarra et al. 2003
Cyanobacteria	Limnospira maxima	nutritional supplements	Priyadarshani & Rath 2012
	Arthrospira platensis	Human nutritional supplements	Colla et al. 2007
Bacillariophyta	Odontella aurita	Fatty acids	Pasquet et al. 2014
	Phaedactylum tricornutum		Yongmanitchai & Ward 1991
Euglenophyta	Euglena gracilis	Vitamin E	Takeyama et al. 1997
Rhodophyta	Porphyridium cruentum	Polysaccharides	Balti et al. 2018

Although many microalgae and cyanobacteria are cultured, only four species have been the focus for biotechnological application through the last decades: Arthrospira platensis Gomont, 1892 (commercially known and marketed as Spirulina), Chlorella vulgaris Beyerinck, 1890), Dunaliella salina Teodoresco, 1905, and Haematococcus pluvialis Flotow, 1844 (Mobin & Alam 2017). More recent studies expanded the attention to other microalgae with potential for biofuel production, but large scale cultivation is still rare. Among the four commercially important species mentioned above, Arthrospira platensis is a Cyanobacteria, but all the other are chlorophytes. Chlorophytes have a thick cell wall that exhibits a wide variety of chemical composition and morphology within the group (Rashidi & Trindade 2018). In fact, a great variability has been reported for the cell wall composition and structure among chlorophyte genera, species and even among lineages or the life stage of the cell (Domozych et al. 2012).

As mentioned before in this review, little is known about the cell wall structure for many species of microalgae (Scholz et al. 2014) and generalizations are frequently made based on few studies considering a very small number of species. This is frequently a problem, since the compounds of interest are mostly found within the cells (Baudelet et al. 2017) and the cell wall can be acting as a barrier to access these products (Kim et al. 2016). Taxonomy, although being a strong tool, is not enough to appropriately deduce the composition of the algae wall in order to reduce costs and time for development of rupture processes for them (Baudelet et al. 2017). In this context, knowledge of the composition and architecture of algal cell coverings is essential to optimize the extraction and recovery of the compounds of commercial interest (Dixon & Wilken 2018).

The lack of knowledge on algal cell coverings contributes with the difficulty to disrupt algal cells to extract compounds, which is one of the biggest obstacles to the industrial use of microalgae on a large scale (Wu et al. 2017). Several methods of disrupting microalgae cells were developed, and their applications depend on the characteristics of the cell and on which compounds are of interest (Dixon & Wilken 2018). These methods can be mechanical or non-mechanical. Mechanical methods include treatments with high pressure homogenization techniques, high speed homogenization, ultrasound and pulsed electric field (Aarthy et al. 2018). Non-mechanical methods can be thermal (microwave, autoclave or freezing), chemical (organic solvent, osmotic shock and acid-alkaline reactions) and biological (microbial or enzymatic degradation) (Dixon & Wilken 2018). Despite all these methods, the disruption of algae cell coverings remains a problem since they are often expensive and inefficient. Cell disruption is crucial for the valorization of algal biomass, however, obtaining efficient and economically attractive cell disruption methods and for all species of interest is still a challenge (D’Hondt et al. 2017).

Conclusion

As mentioned in this review, cell coverings generally play important roles in cellular physiology and each type of covering must meet the specific needs of each group. This study tried to organize the knowledge about the several types of coverings of microalgae and cyanobacteria and highlighted how diverse they are. Cell coverings can be intra or extracellular and have a variety of mainly components depending of group and most of them are considered a taxonomic feature due to their group or species-specific morphology. This review allows us to notice that despite all the research cited, very little is known about microalgae coverings, considering the diversity of species. Some few groups, such as diatoms, are in general the most studied group due to their possibilities of use in the areas of bio and nanotechnology. However, algae in general are a very diverse group and have been increasingly studied for different purposes, which requires a better understanding of various aspects of these organisms, especially the morphological ones.

References

• Aarthy A, Kumari S, Turkar P, Subramanian S. 2018. An insight on algal cell disruption for biodiesel production. Asian Journal of Pharmaceutical and Clinical Research 11: 21-26.
• Abo-shady AM, Mohamed YA, Lasheen T. 1993. Chemical composition of the cell wall in some green algae species. Biologia Plantarum 35: 629-632.
• Aiad I, Riya MA, Tawfik SM, Abousehly MA. 2016. Synthesis, surface properties and biological activity of N, N, N-tris(hydroxymethyl)-2-oxo-2-(2-(2-(alkanoyloxy) ethoxy) ethoxy) ethanaminium chloride surfactants. Egyptian Journal of Petroleum 25: 299-307.
• Arora M, Anil AC, Leliaert F, Delany J, Mesbahi E. 2013. Tetraselmis indica (Chlorodendrophyceae, Chlorophyta), a new species isolated from saltpans in Goa, India. European Journal of Phycology 48: 61-78.
• Baldan B, Andolfo P, Navazio L, Tolomio C, Mariani P. 2001. Cellulose in algal cell wall: an “in situ” localization. European Journal of Histochemistry 45: 51-56.
• Balti R, Le-Balc’h R, Brodu N, et al 2018.Concentration and purification of Porphyridium cruentum exopolysaccharides by membrane filtration at various cross-flow velocities. Process Biochemistry 74: 175-184.
• Bandarra MN, Pereira PA, Batista I, Vilela MH. 2003. Fatty acids, sterols and α-tocopherol in Isochrysis galbana Journal of Food Lipids 10: 25-34.
• Baudelet PH, Ricochon G, Linder M, Muniglia L. 2017. A new insight into cell walls of Chlorophyta. Algal Research 25: 333-371.
• Becker B, Marin B, Melkonian M. 1994. Structure, composition and biogenesis of prasynophyte cell coverings. Protoplasma: 181: 233-244.
• Belcher JH. 1969. A morphological study of the phytoflagellate Chrysococcus Rufescens Klebs in culture. British Phycological Journal 4: 105-117.
• Bisalputra T, Weier W. 1963. The cell wall of Scenedesmus quadricauda American Journal of Botany 50: 1011-1019.
• Bown PR, Lees JA, Young JR. 2004. Calcareous nannoplankton evolution and diversity through time. In: Thierstein HR, Young JR. (eds.) Coccolithophores: From Molecular processes to Global Impact. Berlin, Springer. p. 481-508.
• Braarud T, Deflandre G, Halldal P, Kamptner E. 1995. Terminology, nomenclature, and systematics of the Coccolithophoridae. Micropaleontology 1: 157-159.
• Brec LC, Kugrens P, Lee RE. 1999. A revised classification of Cryptophyta. Botanical Journal of the Linnean Society 131: 131-151.
• Brett SJ, Perasso L, Wetherbee R. 1994. Structure and development of the cryptomonad Periplast: a review. Protoplasma 181: 106-122.
• Brett SJ, Wetherbee R. 1986. A Comparative Study of Periplast Structure in Cryptomonas cryophila and C. ovata Protoplasma 131: 23-31.
• Brett SJ, Wetherbee R. 1996a. Periplast development in Cryptophyceae II. Development of the inner periplast component in Rhinomonas pauea, Proteomonas sulcata [haplomorph], Rhodomonas baltica and Cryptomonas ovata Protoplasma 192: 40-48.
• Brett SJ, Wetherbee R. 1996b. Periplast development in Cryptophyceae III. Development of crystalline surface plates in Falcomonas daucoides, Proteomonas sulcata [haplomorph], and Komma caudata Protoplasma 192: 49-56.
• Brosnan S, Brown PJP, Farmer MA, Triemer RE. 2005. Morphological separation of the euglenoid genera Trachelomonas and Strombomonas (Euglenophyta) based on lorica development and posterior strip reduction. Journal of Phycology 41: 590-605.
• Cavalier-Smith T. 2016. Higher classification and phylogeny of Euglenozoa. European Journal of Protistology 56: 250-276.
• Cavalier-Smith T. 2017. Euglenoid pellicle morphogenesis and evolution in light of comparative ultrastructure and trypanosomatid biology: Semi-conservative microtubule strip duplication, strip shaping and transformation. European Journal of Protistology 61: 137-179.
• Chan WS, Kwok ACM, Wong JTY. 2019. Knockdown of Dinoflagellate Cellulose Synthase CesA1 Resulted in Malformed Intracellular Cellulosic Thecal Plates and Severely Impeded Cyst-to-Swarmer Transition. Frontiers in Microbiology: 10: 546. doi: 10.3389/fmicb.2019.00546
  » https://doi.org/10.3389/fmicb.2019.00546
• Chrétiennot-Dinet MJ, Desreumaux N, Vignes-Lebbe R. 2014. An interactive key to the Chrysochromulina species (Haptophyta) described in the literature. PhytoKeys 34: 47-60.
• Colla LM, Reinehr CO, Reichert C, Costa JAC. 2007. Production of biomass and nutraceutical compounds by Spirulina platensis under different temperature and nitrogen regimes. Bioresource Technology 98: 1489-1493.
• Conforti V. 2010. Ultrastructure of the lorica of species (Euglenophyta) from New Jersey, USA. Algological Studies 135:15- 40.
• D’Hondt E, Martin-Juarez J, Bolado S, Kasperoviciene J, Kor-eiviene J, Sulcius S, Bastiaens L. 2017. Cell disruption technologies. In: Muñoz R, Gonzalez-Fernandez C. (eds.) Microalgae-based biofuels and bioproducts. Cambridge, UK, Woodhead Publishing. p. 133-154.
• Daugbjerg N. 2000. Pyramimonas tychotreta sp. nov. (Prasinophyceae), a new marine species from Antarctica: light and electron microscopy of the motile stage and notes on growth rates. Journal of Phycology 36: 160-71.
• Dixon C, Wilken LR. 2018. Green microalgae biomolecule separations and recovery. Bioresources and Bioprocessing 5: 1-24.
• Dodge JD, Crawford RM. 1970. A survey of thecal fine structure in the Dinophyceae. Botanical Journal of the Linnean Society 63: 53-67.
• Dodge JD. 1983. Dinoflagellates: Investigation and phylogenetic speculation. British Phycological Journal 18: 335-356.
• Doerrler WT. 2006. Lipid trafficking to the outer membrane of Gram-negative bacteria. Molecular Microbiology 60: 542-552.
• Dolatabadi JEN, La-Guardia M. 2011. Applications of diatoms and silica nanotechnology in biosensing, drug and gene delivery, and formation of complex metal nanostructures. Trends in Analytical Chemistry 30: 1538-1548.
• Domozych DS, Brechka H, Britton A, Toso M. 2011. Cell wall growth and modulation dynamics in a model unicellular green alga - Penium margaritaceum: Live cell labeling with monoclonal antibodies. Journal of Botany 2011: 632165. doi: 10.1155/2011/632165
  » https://doi.org/10.1155/2011/632165
• Domozych DS, Ciancia M, Fangel JU, Mikkelsen MD, Ulvskov P, Willats WGT. 2012. The cell walls of green algae: a journey through evolution and diversity. Frontiers in Plant Science 3: 82. doi: 10.3389/fpls.2012.00082
  » https://doi.org/10.3389/fpls.2012.00082
• Domozych DS, Serfis A, Kiemle SN, Gretz MR. 2007. The structure and biochemistry of charophycean cell walls: I. Pectins of Penium margaritaceum Protoplasma 230: 99-115.
• Domozych DS, Sørensen I, Pettolino FA, Bacic A, Willats WGT. 2010. The cell wall polymers of the charophycean green alga Chara corallina: immunobinding and biochemical screening. International Journal of Plant Sciences 171: 345-361.
• Duangjan K, Wołowski K. 2013. New taxa of loricate euglenoids Strombomonas and Trachelomonas from Thailand. Polish Botanical Journal 58: 337-345.
• Dunlap JR, Walne PL, Preisig HR. 1987. Manganese mineralization in chrysophycean loricas. Phycologya 26: 394-396.
• Eder M, Lütz-Meindl U. 2008. Pectin-like carbohydrates in the green alga Micrasterias characterized by cytochemical analysis and energy filtering TEM. Journal of Microscopy 231: 201-214.
• Eder M, Lütz-Meindl U. 2010. Analyses and localization of pectin-like carbohydrates in cell wall and mucilage of the green alga Netrium digitus Protoplasma 243: 25-38.
• Eder M, Tenhaken R, Driouich A, Lütz-Meindl U. 2008. Occurrence and characterization of arabinogalactan-like proteins and hemicelluloses in Micrasterias (Streptophyta). Journal of Phycology 44: 1221-1234.
• Eikrem W, Medlin LK, Henderiks J, Rokitta SD, Rost I. 2017. Haptophyta. In: Archibald JM, Simpson AGB, Slamovits CH. (eds.) Handbook of the Protists, 2nd edn. German, Springer
• Estevez JM, Fernández PV, Kasulin L, Dupree P, Ciancia M. 2009. Chemical and in situ characterization of macromolecular components of the cell walls from the green seaweed Codium fragile Glycobiology 19: 212-228.
• Faber WW, Preisig HR. 1994. Calcified structures and calcification in protists. Protoplasma 181: 78-105.
• Faust MA. 1974. Structure of the periplast of Cryptomonas ovata var. palustris. Journal of Phycology 10: 121-124.
• Ford CW, Percival E. 1965. The carbohydrates of Phaeodactylum tricornutum. Part II. A sulphated glucuronomannan. Journal of the Chemical Society 129: 7042-7046.
• Fry SC. 2003. The plant cell wall. Oxford, Routledge & CRC Press.
• Gantt E. 1971. Micromorphology of the periplast of Chroomonas sp. (Cryptophyceae). Journal of Phycology 7: 177-184.
• Gaysina LA, Saraf A, Singh P. 2019. Cyanobacteria in Diverse Habitats. In: Mishra AK, Tiwari DN, Rai AN. (eds.) Cyanobacteria. Amsterdam, Elsevier. p. 1-28.
• Gigova L, Marinova G. 2016. Significance of microalgae-grounds and areas. Genet Plant Physiol 6: 85-100.
• Gómez F. 2007. Gymnodinioid Dinoflagellates (Gymnodiniales, Dinophyceae) in the Open Pacific Ocean. Algae 22: 273-286.
• Goodenough UW, Heuser JE. 1985. The Chlamydomonas cell wall and its constituent glycoproteins analyzed by the quick-freeze, deep-etch technique. The Journal of Cell Biology 101: 1550-1568.
• Gügi B, Costaouec T, Burel C, Lerouge P, Helbert W, Bardor M. 2015. Diatom-specific oligosaccharide and polysaccharide structures help to unravel biosynthetic capabilities in diatoms. Marine Drugs 13: 5993-6018 .
• Harðardóttir S, Lundholm N, Moestrup Ø, Nielsen TG. 2014. Description of Pyramimonas diskoicola sp. nov. and the importance of the flagellate Pyramimonas (Prasinophyceae) in Greenland sea ice during the winter-spring transition. Polar Biology 37:1479-1494.
• Hausmann K. 1978. Extrusive organelles in protists. International Review of Cytology 52: 197-276.
• Hausmann K. 1979. The Function of the Periplast of the Cryptophyceae during the Discharge of Ejectisomes. Archieves Protistenk 122: 222-225.
• He X, Dai J, Wu Q. 2016. Identification of sporopollenin as the outer layer of cell wall in microalga Chlorella protothecoides Frontiers of Microbiology 7: 1047. doi: 10.3389/fmicb.2016.01047.
  » https://doi.org/10.3389/fmicb.2016.01047
• Hecky RE, Mopper K, Kilham P, Degens ET. 1973. Amino acid and sugar composition of diatom cell walls. Marine Biology 19: 323-31.
• Herburger K, Holzinger A. 2015. Localization and quantification of callose in the streptophyte green algae Zygnema and Klebsormidium: Correlation with desiccation tolerance. Plant Cell Physiology 56: 2259-2270.
• Herth W, Kuppel A, Schnepf E. 1977. Chitinous fibrils in the lorica of the Flagellate chrysophyte Poteriochromonas stipitata (syn. Ochromonas malhamensis). The Journal of Cell Biology 73: 311-321.
• Herth W, Zugenmaier P. 1979. The Lorica of Dinobryon Journal of Ultrastructure Research 69: 262-272.
• Hiremath PS, Bannigidad P. 2011. Automated Gram-staining characterisation of bacterial cells using colour and cell wall properties. International Journal of Biomedical Engineering and Technology 7: 257-265.
• Hoef-Emden K, Melkonian M. 2003. Revision of the Genus Cryptomonas (Cryptophyceae): A combination of molecular phylogeny and morphology provides insights into a long-hidden dimorphism. Protist 154: 371-409.
• Höhfeld I, Otten J, Melkonian M. 1988. Contractile eucaryotic flagella: Centrin is involved. Protoplasma 147: 16-24.
• Hoiczyk E, Hansel A. 2000. Cyanobacterial cell walls: News from an unusual prokaryotic envelope. Journal of Bacteriology 182: 1191-1199.
• Hoson T. 2002. Physiological functions of plant cell coverings. Journal of Plant Research 115: 277-282.
• Houdan AC, Marie D, Not F, Saez AG, Young JR, Probert I. 2004. Holococcolithophore-heterococcolithophore (Haptophyta) life cycles: Flow cytometric analysis of relative ploidy levels. Systematics and Biodiversity 4: 453-465.
• Imam SH. 1985. The Chlamydomonas cell wall: characterization of the wall framework. The Journal of Cell Biology 101: 1599-1607.
• Irazoki O, Hernandez SB, Cava F. 2019. Peptidoglycan muropeptides: Release, perception and functions as signaling molecules. Frontiers in Microbiology 10: 500. doi: 10.3389/fmicb.2019.00500
  » https://doi.org/10.3389/fmicb.2019.00500
• Jamali AA, Akbari F, Ghorakhlu MM, La-Guardia M, Khosroushahi AY. 2012. Applications of Diatoms as Potential Microalgae in Nanobiotechnology. BioImpacts 2: 83-89.
• Jönsson M, Allahgholi L, Sardari RRR, Hreggviðsson GO, Karlsson EN. 2020. Extraction and Modification of Macroalgal Polysaccharides for Current and Next-Generation Applications. Molecules 25: 930. doi: 10.3390/molecules25040930
  » https://doi.org/10.3390/molecules25040930
• Jordan RW, Chamberlain AHL. 1997. Biodiversity among haptophyte algae. Biodiversity and Conservation 6: 131-152.
• Jurgens UJ, Weckesser J. 1985. The fine structure and chemical composition of the cell wall and sheath layers of cyanobacteria. Annales de L' Institut Pasteur Microbiologie 136: 41-44.
• Kapustin DA. 2019. Nomenclatural notes on triporous taxa of Chrysococcus G.A. Klebs (Chrysophyceae). Phytotaxa 387: 71-72.
• Karim AGA, Round FE. 1967. Microfibrils in the lorica of the freshwater alga Dinobryon The New Phytologist 66: 409-412.
• Kim DY, Vijayan D, Praveenkumar R, et al 2016. Cell-wall disruption and lipid/astaxanthin extraction from microalgae: Chlorella and Haematococcus Bioresource Technology 199: 300-310.
• Kristiansen J, Škaloud P. 2017. Chrysophyta. In: Archibald JM, Simpson AGB, Slamovits CH. (eds.) Handbook of the Protists. German, Springer International Publishing AGJM. p. 331-366.
• Kristiansen J. 1986. The ultrastructural bases of chrysophyte systematics and phylogeny. Critical Reviews in Plant Sciences 4: 149-211.
• Kristiansen J. 2002. The genus Mallomonas (Synurophyceae) - a taxonomic survey based on the ultrastructure of silica scales and bristles. Opera Botanica 139: 5-218 .
• Kristiansen J. 2008. Dispersal and biogeography of silica-scaled chrysophytes. Biodiversity and Conservation 17: 419-426.
• Kröger N, Poulsen N. 2008. Diatoms-From cell wall biogenesis to nanotechnology. Annual Reviews of Genetics 42: 83-107.
• Kröger N, Deutzmann R, Sumper M. 2001. Silica-precipitating peptides from diatoms. The chemical structure of silaffin-A from Cylindrotheca fusiformis Journal of Biological Chemistry 276: 26066-70.
• Kröger N, Deutzmann R, Bergsdorf C, Sumper M. 2000. Species-specific polyamines from diatoms control silica morphology. Proceedings of the National Academy of Sciences 97: 14133-38.
• Kröger N, Deutzmann R, Sumper M. 1999. Polycationic peptides from diatom biosilica that direct silica nanosphere formation. Science 286: 1129-1132.
• Kröger N, Lehmann G, Rachel R, Sumper M. 1997. Characterization of a 200-kDa diatom protein that is specifically associated with a silica-based substructure of the cell wall. European Journal of Biochemistry 250: 99-105.
• Kröger N, Bergsdorf C, Sumper M. 1996. Frustulins: domain conservation in a protein family associated with diatom cell walls. European Journal of Biochemistry 239: 259-64.
• Kröger N, Bergsdorf C, Sumper M. 1994. A new calcium-binding glycoprotein family constitutes a major diatom cell wall component. The EMBO Journal 13: 4676-83.
• Kugrens P, Lee RE. 1987. An ultrastructural survey of cryptomonad periplasts using Quick-freezing freeze fracture techniques. Journal of Phycology 23: 365-376.
• Kugrens P, Lee RE. 1991. Organization of Cryptomonads. In: Patterson OJ, Larsen J. (eds) The biology of free-living heterotrophic flagellates. The Systematics Association Special Volume 45: 219-233.
• Leadbeater BSC, Barker DAN. 1995. Biomineralization and scale production in the Chrysophyta. In: Sandgren CD, Smol JP, Kristiansen J. (eds.) Chrysophyte algae: ecology, phylogeny and development. Cambridge, Cambridge University Press. p. 141-164.
• Leander BS, Esson HJ, Breglia SA. 2007. Macroevolution of complex cytoskeletal systems in euglenids. BioEssays 29: 987-1000.
• Leander BS, Witek RP, Farmer MA. 2001. Trends in the evolution of the euglenid pellicle. Evolution 55: 2215-2235.
• Lee RE. 2008. Phycology. 4th edn. Cambridge, Cambridge University Press
• Leedale GF. 1964. Pellicle structure in Euglena British Phycological Bulletin 2: 291-306.
• Leedale GF. 1975. Envelope formation and structure in the euglenoid genus Trachelomonas British Phycological Society 10:17-41.
• Liu H, Aris-Brosou S, Probert I, Vargas C. 2010. A time line of the environmental genetics of the haptophytes. Molecular Biology and Evolution 27: 161-176.
• Maher S, Kumeria T, Aw MS, Losic D. 2018 . Diatom silica for biomedical applications: Recent progress and advances. Advanced Healthcare Materials 7: 1-19.
• Maldonado RF, Sá-Correia I, Valvano MA. 2016. Lipopolysaccharide modification in gram-negative bacteria during chronic infection. FEMS Microbiology Reviews 40: 480-493.
• Manton I. 1966. Observations on scale production in Prymnesium parvum Journal of Cell Science 1: 375-380.
• Metzger P, Largeau C. 2005. Botryococcus braunii: A rich source for hydrocarbons and related ether lipids. Applied Microbiology and Biotechnology 66: 486-496.
• Mobin S, Alam F. 2017. Some promising microalgal species for commercial applications: a review. Energy Procedia 110: 510-517.
• Moestrup O, Walne PL. 1979. Studies on scale morphogenesis in the Golgi apparatus of Pyramimonas tetrarhynchus (Prasinophyceae). Journal of Cell Science 36: 437- 459.
• Monteiro FM, Bach LT, Brownlee C, et al 2016. Why marine phytoplankton calcify. Science Advances 2: e1501822. doi: 10.1126 / sciadv.1501822
  » https://doi.org/10.1126 / sciadv.1501822
• Morrill LC, Loeblich AR. 1983. Ultrastructure of the dinoflagellate amphiesma. International Review of Cytology 82: 151-180.
• Morrill LC. 1984. Ecdysis and the location of the plasma membrane in the dinoflagellate Heterocapsa niei Protoplasma 119: 8-20.
• Müller MN. 2019. On the genesis and function of coccolithophore calcification. Frontiers in Marine Science 6: 49. doi: 10.3389/fmars.2019.00049
  » https://doi.org/10.3389/fmars.2019.00049
• Nakajima T, Volcani BE. 1969. 3,4 Dihydroxyproline: a new aminoacid in diatom cell walls. Science 164: 1400-1401.
• Němcová Y, Pichrtová M. 2012. Shape dynamics of silica scales (Chrysophyceae, Stramenopiles) associated with pH. Fottea, Olomouc 12: 281-291.
• Němcová Y. 2003. Detection of cell wall structural polysaccharides by cellulase-gold and chitinase-gold complexes. Czech Phycology Olomouc 3:31-36.
• Neustupa J, Řezáčová-Škaloudová M, Němcová Y. 2010. Shape variation of the silica-scales of Mallomonas kalinae (Mallomonadales, Synurophyceae) in relation to their position on the cell body. Nova Hedwigia 136: 33-41.
• Nikaido H. 2003. Molecular Basis of Bacterial Outer Membrane Permeability Revisited. Microbiology and Molecular Biology Reviews 67: 593- 656.
• Okuda K, Sekida S. 2007. Cellulose-synthesizing complexes of a dinoflagellate and other unique algae. In: Brown Jr. RM, Saxena IM. (eds.) Cellulose: Molecular and Structural Biology. German, Spring. p. 199-215.
• Okuda K. 2002. Structure and phylogeny of cell coverings. Journal of Plant Research 115: 283-288.
• Paasche E. 2002. A review of the coccolithophorid Emiliania huxleyi (Prymnesiophyceae), with particular reference to growth, coccolith formation, and calcification-photosynthesis interactions. Phycologia 40: 503-529.
• Parker BC. 1964. The structure and chemical composition of cell walls of three chlorophycean algae. Phycologia 4: 63-74.
• Pasquet V, Ulmann L, Mimouni V, et al 2014. Fatty acids profile and temperature in the cultured marine diatom Odontella aurita Journal of Applied Phycology 26: 2265-2271.
• Peck RK. 2010. Structure of loricae and stalks of several bacterivorous Chrysomonads (Chrysophyceae): taxonomical importance and possible ecological significance. Protist 161: 148-159.
• Perasso L, Ludwig M, Wheterbee R. 1997. The surface periplast component of the protist Komma caudata (Cryptophyceae) self-assembles from a secreted high-molecular-mass polypeptide. Protoplasma 200: 186-197.
• Pereira H, Silva J, Santos T, et al 2019. Nutritional potential and toxicological evaluation of Tetraselmis sp. CTP4 microalgal biomass produced in industrial photobioreactors. Molecules 24: 3192. doi: 10.3390/molecules24173192
  » https://doi.org/10.3390/molecules24173192
• Peterson PK, Quie PG. 1981. Bacterial surface components and the Pathogenesis of infectious diseases. Annual Review of Medicine 32: 29-43.
• Piro G, Lenucci M, Dalessandro G, et al 2000. Ultrastructure, chemical composition and biosynthesis of the cell wall in Koliella antarctica (Klebsormidiales, Chlorophyta). European Journal of Phycology 35: 331-337.
• Poniewozik M. 2017. Element composition of Trachelomonas envelopes (Euglenophyta). Polish Botanical Journal 62: 77-85.
• Popper ZA, Fry SC. 2003. Primary cell wall composition of bryophytes and charophytes. Annals of Botany 91: 1-12.
• Pozdnyakov I, Skarlato S. 2012. Dinoflagellate amphiesma at different stages of the life cycle. Protistology 7: 108-115.
• Priyadarshani I, Rath B. 2012. Commercial and industrial applications of micro algae- a review. Journal of Algal Biomass Utilization 3: 89-100.
• Raja R, Hemaiswarya S, Rengasamy R. 2007. Exploitation of Dunaliella for β-carotene production. Applied Microbiology and Biotechnology 4:517-523.
• Raja R, Shanmugam H, Ganesan V, Carvalho IS. 2014. Biomass from microalgae: an overview. Oceanography 2: 1-7.
• Rashidi B, Trindade LM. 2018 . Detailed biochemical and morphologic characteristics of the green microalgaNeochloris oleoabundanscell wall. Algal Research 35: 152-159.
• Reimann BEF, Lewin JC, Volcani BE. 1965. Studies on the biochemistry and fine structure of silica shell formation in diatoms. I. The structure of the cell wall of Cylindrotheca fusiformis Reimann and Lewin. Journal of Cell Biology 24: 39-55.
• Resch CM, Gibson J. 1983. Isolation of the carotenoid-containing cell wall of three unicellular cyanobacteria. Journal of Bacteriology 155: 345-350.
• Řezáčová-Škaloudová M, Neustupa J, Němcová Y. 2010. Effect of temperature on the variability of silicate structures in Mallomonas kalinae and Synura curtispina (Synurophyceae). Nova Hedwigia. Beihefte 136:55-70.
• Rudolf M, Tetik N, Ramos-León F, et al 2015. The peptidoglycan-binding protein SjcF1 influences septal junction function and channel formation in the filamentous cyanobacterium Anabaena MBio 6:4. doi: 10.1128 / mBio.00376-15
  » https://doi.org/10.1128 / mBio.00376-15
• Saad A, Atia A. 2014. Review on Freshwater Blue-Green Algae (Cyanobacteria): Occurrence, Classification and Toxicology. Biosciences Biotechnology Research 11: 1319-1325.
• Safi C, Camy S, Frances C, et al 2014. Extraction of lipids and pigments of Chlorella vulgaris by supercritical carbon dioxide: influence of bead milling on extraction performance. Journal of Applied Phychology: 26: 1711-1718 .
• Saldarriaga JF, ‘Max’ Taylor FJR. 2017. Dinoflagellata. In: Archibald JM, Simpson AGB, Slamovits CH. (eds.) Handbook of the Protists. Cham, Springer. p. 625-678.
• Sanders WB. 2001. Lichens: The Interface between Mycology and Plant Morphology. BioScience 51: 1025-1035.
• Sandgren CD, Hall SA, Barlow SB. 1996. Siliceous scale roduction in Chrysophyte and Synurophyte algae. I. Effects of silica-limited growth on cell silica content, scale morphology, and the construction of the scale layer of Synura petersenii Journal of Phycology 33: 675-692.
• Scholz MJ, Weiss TL, Jinkerson RE, at al 2014. Ultrastructure and Composition of the Nannochloropsis gaditana Cell Wall. Eukaryotic Cell 13: 1450-1464.
• Schütt F. 1895. Die peridineen der plankton-expedition. Ergebnisse der Plankton-Expedition der Humboldt-Stiftung 4: 1-170.
• Schwelitz FD, Evans WR, Mollenhauer HH, Dilley RA. 1970. The Fine Structure of the Pellicle of Euglena gracilis as revealed by Freeze-etching. Protoplasma 69: 341-349.
• Sekida S, Horiguchi T, Okuda K. 2001. Development of the cell covering in the dinoflagellate Scrippsiella hexapraecingula (Peridiniales, Dinophyceae). Phycological Research 49: 163-176.
• Sekida S, Horiguchi T, Okuda K. 2004. Development of thecal plates and pellicle in the dinoflagellate Scrippsiella hexapraecingula (Peridiniales, Dinophyceae) elucidated by changes in stainability of the associated membranes. European Journal of Phycology 39: 105-114.
• Shah MMR, Liang Y, Cheng JJ, Daroch M. 2016. Astaxanthin-producing green microalga Haematococcus pluvialis: from single cell to high value commercial products. Frontiers in Plant Science 7: 531. doi: 10.3389/fpls.2016.00531
  » https://doi.org/10.3389/fpls.2016.00531
• Sikes CS, Roer RD, Wilbur KM. 1980. Photosynthesis and coccolith formation: Inorganic carbon sources and net inorganic reaction of deposition. Limnology and Oceanography 252: 48-261.
• Silhavy TJ, Kahne D, Walker S. 2010. The Bacterial Cell Envelope. Cold Spring Harbor Perspectives in Biology 2: a000414. doi: 10.1101 / cshperspect.a000414
  » https://doi.org/10.1101 / cshperspect.a000414
• Siver PA .1989. The distribution of scaled chrysophytes along a pH gradient. Canadian Journal of Botany 67: 2120-2130.
• Škaloud P, Kristiansen J, Škaloudová M. 2013. Developments in the taxonomy of silica-scaled chrysophytes - from morphological and ultrastructural to molecular approaches. Nordic Journal of Botany 31: 385-402.
• Škaloud P, Kynčlová A, Benada O, Kofroňová O, Škaloudová M. 2012. Toward a revision of the genus Synura, section Petersenianae (Synurophyceae, Heterokontophyta): morphological characterization of six pseudo-cryptic species. Phycologia 51: 303-329.
• Sommer JR. 1965. The ultrastructure of the pellicle complex of Euglena gracilis. The Journal of Cell Biology 24: 253-257.
• Sørensen I, Pettolino FA, Bacic A, et al 2011. The charophycean green algae provide insights into the early origins of plant cell walls. Plant Journal 68: 201-211.
• Strother PK, Taylor WA, Schootbrugge B, Leander BS, Wellman CH. 2019. Pellicle ultrastructure demonstrates that Moyeria is a fossil euglenid. Palynology 44: 461-471.
• Swift E, Remsen CC. 1970. The cell wall of Pyrocystis spp. (Dinococcales). Journal of Phycology 6: 79-86.
• Takeyama H, Kanamaru A, Yoshino Y, Kakuta H, Kawamura Y, Matsunaga T. 1997. Production of Antioxidant Vitamins, β-Carotene, Vitamin C, and Vitamin E, by Two-Step Culture of Euglena gracilis Z. Biotechnology and Bioengineering 53: 185-190.
• Taylor AR, Brownlee C, Wheeler G. 2016. Coccolithophore cell biology: Chalking Up progress. Annuals Review of Marine Science 9: 283-310.
• Terraciano M, Stefano L, Rea I. 2018 . Diatoms green nanotechnology for biosilica-based drug delivery systems. Pharmaceutics 10: 242. doi: 10.3390/pharmaceutics10040242
  » https://doi.org/10.3390/pharmaceutics10040242
• Tesson B, Hildebrand M. 2010. Extensive and intimate association of the cytoskeleton with forming silica in diatoms: Control over patterning on the meso- and micro-scale. PLOS ONE 5: e14300. doi: 10.1371/journal.pone.0014300.
  » https://doi.org/10.1371/journal.pone.0014300
• Tommasi G, Gielis J, Rogato A. 2017. Diatom frustule morphogenesis and function: a multidisciplinary survey. Marine Genomics 35: 1-18 .
• Vargas C, Aubry MP, Probert I,Young J. 2007. Origin and evolution of coccolithophores: from coastal hunters to oceanic farmers. In: Falkowski PG, Knoll AH. (eds.) Evolution of Primary Producers in the Sea. New York, Elsevier. p. 51- 285.
• Vismara R, Barsanti L, Lupetti P, et al 2000. Ultrastructure of the pellicle of Euglena gracilis Tissue and Cell 32: 451-456.
• Vollmer W, Blanot D, de Pedro MA. 2008. Peptidoglycan structure and architecture. FEMS Microbiology Reviews 32: 149-167.
• Walker JM, Marzec B, Lee RBY, et al 2018 . Polymorph Selectivity of Coccolith-Associated Polysaccharides from Gephyrocapsa Oceanica on Calcium Carbonate Formation in Vitro. Advanced Functional Materials 29: 1807168. doi: 10.1002/adfm.201807168
  » https://doi.org/10.1002/adfm.201807168
• Wang DZ, Dong HP, Li C, Xie ZX, Lin L, Hong HS. 2011. Identification and characterization of cell wall proteins of a toxic dinoflagellate Alexandrium catenella using 2-D DIGE and MALDI TOF-TOF mass spectrometry. Evidence-Based Complementary and Alternative Medicine: eCAM 2011: 984080. doi: 10.1155/2011/984080
  » https://doi.org/10.1155/2011/984080
• Wee JL. 1997. Scale biogenesis in synurophycean protists: phylogenetic implications. Critical Reviews in Plant Sciences 16: 497-534.
• Wiltshire KH, Boersma M, Möller A, Buhtz H. 2000. Extraction of pigments and fatty acids from the green alga Scenedesmus obliquus (Chlorophyceae). Aquatic Ecology 34: 119-126.
• Wooldridge SA. 2013. Breakdown of the coral-algae symbiosis: Towards formalizing a linkage between warm-water, bleaching thresholds and the growth rate of the intracellular zooxanthellae. Biogeosciences 10: 1647-1658.
• Wu C, Xiao Y, Lin W, et al 2017. Aqueous enzymatic process for cell wall degradation and lipid extraction from Nannochloropsis sp. Bioresource Technology 223: 312-316.
• Wurdack ME. 1923. Chemical composition of the walls of certain algae. The Ohio Journal of Science 23: 181-191.
• Yamada T, Sakaguchi K. 1982. Comparative studies in Chlorella cell walls: induction of protoplast formation. Archives of Microbiology 132: 10-13.
• Yongmanitchai W, Ward P. 1991. Growth of and omega-3 fatty acid production by Phaeodactylum tricornutum under different culture conditions. Applied and Environmental Microbiology 57: 419-425.
• Yoshimi A, Miyazawa K, Abe K. 2017. Function and biosynthesis of cell wall -1,3-glucan in fungi. Journal of Fungi 3: 63. doi: 10.3390/jof3040063
  » https://doi.org/10.3390/jof3040063
• Young JR, Henriksen K. 2003. Biomineralization within vesicles: The calcite of coccoliths. Reviews in Mineralogy and Geochemistry 54: 189-215.
• Young JR. 1994. Variation in Emiliania huxleyi coccolith morphology in samples from the Norwegian EHUX experiment 1992. Sarsia 79: 417-425.
• Zancan S, Trevisan R, Maurizio GP. 2006. Soil algae composition under different agro-ecosystems in North-Eastern Italy. Agriculture, Ecosystems and Environment 112: 1-12.
• Zhang GE, Meredith TC, Kahne D.2013. On the essentiality of lipopolysaccharide to gram-negative bacteria. Current Opinion in Microbiology 16: 779-785.
• Zhang JY, Lin GM, Xing WY, Zhang CC. 2018. Diversity of growth patterns probed in live cyanobacterial cells using a fluorescent analog of a peptidoglycan precursor. Frontiers in Microbiology 9: 791. doi: 10.3389/fmicb.2018.00791
  » https://doi.org/10.3389/fmicb.2018.00791

Publication Dates

• Publication in this collection
  22 Mar 2021
• Date of issue
  Oct-Dec 2020

History

• Received
  30 June 2020
• Accepted
  06 Sept 2020