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Contribution to the embryology of Potamogeton L. (Alismatales: Potamogetonaceae)
Aquatic Botany 93 (2010) 32–38
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Contribution to the embryology of Potamogeton L. (Alismatales: Potamogetonaceae) Elaine Lopes Pereira Nunes ∗ , Mariana Cortes de Lima, Alessandra Ike Coan 1 , Maria Cecília de Chiara Moc¸o 2 Departamento de Botânica, Setor de Ciências Biológicas, Universidade Federal do Paraná, C. Postal 19031, CEP 81531-980, Curitiba, Paraná, Brazil
a r t i c l e
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Article history: Received 3 November 2009 Received in revised form 24 February 2010 Accepted 4 March 2010 Available online 10 March 2010 Keywords: Anther wall Megagametogenesis Megasporogenesis Microgametogenesis Microsporogenesis Monocotyledon
a b s t r a c t Potamogetonaceae currently comprises four genera, including Potamogeton, a near cosmopolitan genus of about 100 species. The historical circumscription of this family is controversial, resulting in fragmentary information on the morphology and anatomy of its genera. The aim of the present study was to investigate the anther and ovule development in three species of Potamogeton in order to clarify the embryological features of this genus. The majority of embryological characters were constant among the three species studied and are in accordance with previous reports. In addition, some new and complementary characteristics of Potamogeton embryology are highlighted, including: monocotyledonous-type of anther wall development and carpels bearing a single ovule that develops a Polygonum-type of megagametophyte. A fail in the micropylar dyad to divide is confirmed as the cause of megaspore triads. The developmental sequence of the ovule confirms its campylotropous curvature. © 2010 Elsevier B.V. All rights reserved.
1. Introduction Potamogetonaceae is a subcosmopolitan and aquatic family (Haynes and Holm-Nielsen, 2003) currently placed within the early-divergent order Alismatales (APG III, 2009). This family has previously been placed in the Helobiae (Engler, 1904), Potamogetonales (Takhtajan, 1997), and Najadales (Cronquist, 1981), based mainly on morphological characters. The circumscription of genera within Potamogetonaceae has also varied among different authors over time. The family has been combined with members of Cymodoceaceae, Najadaceae, Zannichelliaceae and Zosteraceae (Haynes and Holm-Nielsen, 2003). According to Haynes and Holm-Nielsen (2003), the genera included in Potamogetonaceae are: Groenlandia J. Gay, Potamogeton L. and Stuckenia Börner. However, APG II (2003) has merged Zannichelliaceae in Potamogetonaceae, with the addition of three more genera: Althenia Petit, Lepilaena J.Drum. ex Harv and Zannichellia L. More recently, Lindqvist et al. (2006), using non-coding nuclear and plastid DNA data to determine Potamoge-
∗ Corresponding author. Tel.: +55 41 3361 1620; fax: +55 41 3266 2042. E-mail address: [email protected] (E.L.P. Nunes). 1 Present address: Departamento de Botânica, Instituto de Biociências, Universidade Estadual Paulista “Júlio de Mesquita Filho”, C. Postal 199, CEP 13506-900, Rio Claro, São Paulo, Brazil. 2 Present address: Departamento de Botânica, Instituto de Biociências, Universidade Federal do Rio Grande do Sul, Av. Bento Gonc¸alves, 9500, CEP 91509-900, Porto Alegre, Rio Grande do Sul, Brazil. 0304-3770/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.aquabot.2010.03.006
tonaceae relationships, concluded that Potamogeton, Groenlandia and Stuckenia, plus Zannichellia L., form a strongly supported monophyletic group. Potamogeton is the largest genus of Potamogetonaceae, with about 100 species (Haynes and Holm-Nielsen, 2003). It is near cosmopolitan, herbaceous, and dispersed by seeds, turions or rhizomes (Haynes and Holm-Nielsen, 2003). Because of its high phenotypic plasticity, hybridization, polyploidy and aneuploidy (Hollingsworth et al., 1998; Kaplan, 2002; Wang et al., 2007), Potamogeton has been historically divided into many infrageneric categories (e.g., Ascherson and Graebner, 1907; Hagström, 1916; Fernald, 1932; Ogden, 1943). However, no such categories were adopted in more recent taxonomic work on Potamogeton (Wiegleb and Kaplan, 1998), which only included notes on systematic affinities. Similarly, the systematic treatment of Potamogetonaceae for the Flora of North America (Haynes and Hellquist, 2000) and for the Flora Neotropica (Haynes and Holm-Nielsen, 2003) did not include an infrageneric classification. A review of the literature shows that the embryological features of Potamogetonaceae are unstable, mainly because of the aforementioned differences in circumscription. The embryological characters compiled by Johri et al. (1992) included genera currently assigned to other families, such as Syringodium Kütz. (Cymodoceaceae), Phyllospadix Hook. and Zostera L. (Zosteraceae). The correct correspondence of each embryological character to each of the above genera is therefore not clear, even for Potamogeton.
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A summary of embryological characters known for Potamogeton is as follows: endothecium with fibrous thickenings (Gupta, 1934; Johri et al., 1992); microsporogenesis of the successive type (Furness and Rudall, 1999 based on Stenar, 1925 and Schnarf, 1931); ovule primordium with three zones (Takaso and Bouman, 1984); ovule ortho-campylotropous (Takaso and Bouman, 1984), orthotropous (Davis, 1966; Igersheim et al., 2001), or anatropous (Posluszny and Sattler, 1974; Posluszny, 1981), bitegmic and crassinucelate (Takaso and Bouman, 1984); linear or T-shaped arrangements of megaspore tetrads (Johri et al., 1992) and megagametophyte of the Polygonum-type (Takaso and Bouman, 1984). According to Haynes and Holm-Nielsen (2003), the type of anther wall development is unknown. Because of the confusion caused by successive circumscription changes in Potamogetonaceae, the aim of the present study was to investigate the early development of the anther and carpel, focusing on micro- and mega-sporogenesis and gametogenesis, of three species of Potamogeton in order to gain a better understanding of the embryology of the genus.
2. Materials and methods Inflorescences at successive developmental stages of three species of Potamogeton were collected in lakes and rivers of Brazil. Vouchers were deposited in the Herbário do Departamento de Botânica, Universidade Federal do Paraná, Curitiba, Paraná, Brazil (UPCB): Potamogeton illinoensis Morong (UPCB 60501), Potamogeton polygonus Cham. & Schltdl (UPCB 61311), and Potamogeton pusillus L. (UPCB 60500). For light microscope (LM) examination, the material was fixed in 1% glutaraldehyde and 4% formaldehyde in 0.1 M phosphate buffer, pH 7.2 (McDowell and Trump, 1976). Samples were rinsed in phosphate buffer, dehydrated through a graded ethanol series and embedded in Leica historesin. Sections (2–5 m) were cut on a Leica RM2145 microtome using glass knives, stained with toluidine blue (O’Brien et al., 1965) and mounted in Permount. Photomicrographs were taken using a Zeiss Axiolab with a digital camera. The following stains were performed on resin-embedded semi-thin sections: iodine potassium iodide solution for starch (Johansen, 1940), Coomassie Brilliant Blue for protein (Southworth, 1973), Ruthenium Red for pectin (Jensen, 1962), Calcofluor White MR2 for cellulose (Feder and O’Brien, 1968) and Aniline Blue for callose (modified from Arens, 1949). Callose and cellulose epifluorescence was observed using a Zeiss Axiophot microscope with a DAPI filter set. Fixed anthers were stained for lipids with Sudan III (O’Brien and McCully, 1981) and for lignin with acid phloroglucin (Sass, 1951). Fixed mature anthers were macerated for 4 h at 60 ◦ C in acetic acid and hydrogen peroxide (1H2 O2 (30%):4 distillate water:5 glacial acetic acid; modified from Ruzin, 1999) and mounted in glycerin jelly to allow observation of the three-dimensional structure of endothecium.
3. Results 3.1. Anther development The flowers of Potamogeton are composed of four tepals, four stamens and four free carpels. In longitudinal sections of the flower, each anther primordium arises as a small group of undifferentiated cells above each tepal primordium (Fig. 1A and B). Protodermal cells give rise to the epidermis through anticlinal divisions (Fig. 1C and D). Archesporial cells are produced by the ground meristem (Fig. 1B and C).
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The development of the anther wall is initiated by the differentiation of the archesporial cells into sporogenous and primary parietal cells (Fig. 1D – arrow). The primary parietal cells undergo periclinal division to produce two secondary parietal layers (Fig. 1E – arrow). The outer layer develops directly into the endothecium, and the inner layer undergoes periclinal divisions to originate two to three ephemerous middle layers and the tapetum (Fig. 1F – arrow and G). The sporogenous cells differentiate into microsporocytes soon after some mitotic divisions (Fig. 1E–G). During meiosis in microsporocytes, the middle layers are compressed against the endothecium and are no longer apparent in the mature anther (Fig. 1H and I). The tapetum is initially a singlelayered tissue with uninucleate cells (Fig. 1F), except in P. polygonus, whose cells become binucleate slightly before they detach one from another (Fig. 1H and I). In P. illinoensis, some cells of the tapetum divide further, giving rise to up to four layers (not shown). The tapetum is one of the plasmodial type, since its cells lose their walls and migrate into the anther locule (Figs. 1H and I and 2A and B). A periplasmodium was observed until the free microspore stage, and disappeared before the onset of microgametogenesis (Fig. 2B). Microsporocytes are rounded (Fig. 1G) and start meiosis concurrent with the detachment of tapetum cells (Fig. 1H and I). The microsporocytes undergo meiosis I as usual (Fig. 1G–I), followed by cytokinesis and dyad formation (Fig. 1I). After meiosis II, a wide range of tetrad shapes was observed, the most common being the tetragonal and decussate (Fig. 2A and B; detailed observations on this issue are discussed by Nunes et al., 2009). Microgametogenesis begins with the formation of a large vacuole that pushes the nucleus towards the cell wall (Fig. 2C) followed by unequal mitotic division of the microspore, resulting in a small generative cell and a large vegetative cell (Fig. 2D). Before shedding, the generative nucleus divides to form two small sperm cells (Fig. 2E). The cytoplasm of the vegetative cell contains numerous starch grains (Fig. 2E). The mature pollen grain has no apertures. The exine is thin and the intine is pectic–cellulosic in nature. The endothecium has large polyhedral cells and Ushaped pectic–cellulosic thickenings by the time of dehiscence (Fig. 2F). Each microsporangium pair coalesces before pollen shedding (Fig. 2F) and dehiscence is longitudinal. 3.2. Ovule development The gynoecium consists of four carpels in a single whorl (Fig. 1A). A single three-zonate ovule primordium is initiated in the ventral wall of each carpel (Fig. 3A). The ovule develops almost horizontally, filling the locule (Fig. 3B and C). Two integuments with two rows of cells each are formed, eventually expanding to three rows, especially next to the chalaza (Fig. 3B–D). As the ovule develops, it gradually bends down to the carpelary base (Fig. 3C–F). This campylotropous curvature is initially brought about by two processes: increased growth of the adjacent region between the chalaza and the integuments, and growth of the carpel, creating space to accommodate the ovule (Fig. 3C–F – marked area). The micropyle is formed by the inner integument alone (Fig. 3G). At the time the megagametophyte is mature, the median region of the ventral wall of the carpel proliferates near the funiculus (Fig. 3H – arrow). This proliferating tissue puts pressure on the funicular region and accentuates the curvature of the ovule. The nucellus divides periclinally, giving rise to four or five parietal layers. The mature ovule is campylotropous, bitegmic and crassinucelate (Fig. 3F–H). The megasporocyte differentiates after the inception of integuments (Fig. 3C). Meiosis I produces a dyad of cells; the micropylar cell is smaller than the chalazal (not shown). Meiosis II proceeds normally in the chalazal cell, producing two megaspores (Fig. 4A). In the micropylar cell, meiosis II is disrupted (Fig. 4A – arrow). This cell soon degenerates, followed by the adjacent megaspore (Fig. 4B
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Fig. 1. Early stages of anther development in P. polygonus (A–F and H–I) and P. illinoensis (G). (A) Longitudinal section (LS) of floral bud showing whorls inception. (B) LS of floral bud with anther primordium in detail. (C) Transversal section (TS) of anther, showing tissues differentiation. (D) Anther primordium showing primary parietal layer (arrow) and sporogenous tissue differentiated, in TS. (E) Periclinal division of the primary parietal layer (arrow), in TS. (F) Anther after a periclinal division of the inner secondary parietal layer (arrow), in TS. (G) Pre-meiotic anther with parietal layers and microsporocytes differentiated, in TS. (H) Anaphase I of meiosis and detachment of binucleate tapetal cells, in TS. (I) Microspore dyads in telophase I. C: carpel; e: epidermis; en: endothecium; MI: microsporocytes; ml: middle layers; pc: procambium; pd: protodermis; S: stamen; sc: sporogenous cells; T: tepal; t: tapetum. Scale bars = 10 m in (A); 20 m in the remaining images.
– arrows). No fully developed megaspore tetrads were observed in any species. Callose was detected only in the transversal walls of the triad. The chalazal-most megaspore is functional (Fig. 4B). Megagametogenesis is brought about by three cycles of mitotic division, resulting in a Polygonum-type megagametophyte (Fig. 4C–E and G–I). When mature, the megagametophyte consists of seven cells: three ephemeral antipodal cells at the chalazal pole (Fig. 4H); the central cell with two polar nuclei that fuse prior to fertilization (Fig. 4H); a single egg cell (Fig. 4G) and two synergids (Fig. 4I) at the micropylar pole. During megasporogenesis and megagametogenesis, the apical epidermal cells and possibly some hypodermal cells of the nucellus divide periclinally to form a nucellar cap (Fig. 4A–F). Neither a hypostase nor an obturator is formed during ovule development.
4. Discussion Anther wall development of Potamogeton is described in detail for the first time in this report. According to the features observed in P. illinoensis, P. polygonus and P. pusillus, anther wall development is of the monocotyledonous-type, in which the inner secondary parietal layer divides to give rise to the middle layers and the tapetum, in accordance with Davis (1966). In the present study, the development of U-shaped pectic–cellulosic thickenings in the endothecial cells was observed at the free microspore stage. Although the presence of thickenings is in agreement with the available data for Potamogeton (Johri et al., 1992; Haynes and Holm-Nielsen, 2003), their nature has remained unclear until now. The type of thickening observed here
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Fig. 2. Late microsporogenesis and microgametogenesis in Potamogeton polygonus (A and C–F) and P. pusillus (B). (A) Microspore tetrads in telophase II and binucleate tapetum cell migrating to the anther locule. (B) Microspore tetrads enclosed by a periplasmodium, in TS. (C) Microgametogenesis onset. (D) Young bicellular pollen grain. (E) Tricellular pollen grain in a mature anther. (F) Anther thecae before dehiscence, in TS. en: endothecium; gc: generative cell; sp: sperm cells; vc: vegetative cell; t: tapetum. Scale bars = 30 m in (J); 10 m in the remaining images.
Fig. 3. Ovule development in Potamogeton polygonus (A, B and D–H) and P. iilinoensis (C). (A) Carpel primordium, in longitudinal section (LS), showing an ovule primordium with three zones. (B) Transversal section (TS) of a carpel with an ovule primordium after the inception of the integuments. (C) Ovule primordium before megasporocyte meiosis in the beginning of its curvature; the development is greater at the interface between chalaza and integuments, in LS. (D–F) Successive stages of ovule curvature due to the greater development of the interface (marked area) between the chalaza and the integuments, in LS. (G) Detail of the micropyle (arrow) of a mature ovule, in LS. (H) LS of ovary showing mature campylotropous ovule, note that the ventral wall of the carpel, adjacent to the the funiculus (arrow), proliferates. ii: inner integument; dw: dorsal wall; n: nucellus; ov: ovule; oi: outer integument; vb: vascular bundle; vw: ventral wall. Scale bars = 50 m in (F) and (H); 20 m in the remaining images.
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Fig. 4. Megasporogenesis and megagametogenesis in Potamogeton polygonus (A–I), ovule in longitudinal sections. (A) Chalazal megaspore dyad and degenerated micropylar dyad (arrow). (B) Functional megaspore at the chalazal pole and degenerated micropylar dyad and adjacent megaspore (arrows). (C) Functional megaspore with cell content at its micropylar region. (D) Immature binucleate megagametophyte (only one nucleus is visible). (E) Four-nucleate megagametophyte (only the two chalazal nuclei are visible). (F) Detail of the micropyle showing the nucellar cap (arrow). (G) Megagametophyte detail showing the egg cell and a synergid partially visible. (H) Megagametophyte detail showing the antipodal cells and the central cell. (I) Polygonum-type megagametophyte. an: antipodals; cc: central cell; ec: egg cell; ii: inner integument; n: nucellus; oi: outer integument; sy: synergid. Scale bars = 50 m in (D) and (E); 10 m in (G) and (H); 20 m in the remaining images.
contrasts with the girdle-type reported by Johri et al. (1992) and Haynes and Holm-Nielsen (2003). In this study, all histochemical staining tests indicated that the endothecial thickenings are definitely composed of pectins and cellulose, but not guaiacyl lignins. However, a negative result of the phloroglucinol–HCl test does not rule out the presence of lignin. Generally, lignins are classified as guaiacyl, guaiacyl–syringyl, or guaiacyl–syringyl–phydroxyphenyl lignins, according to whether they are from gymnosperms, wood angiosperms, or grasses, respectively. A positive result of the phloroglucinol–HCl indicates the presence of coniferaldehyde end-groups in guaiacyl lignins only. However, Lewis and Yamamoto (1990) emphasized that there are exceptions, and that no single analytical method provides conclusive evidence that lignin is present. The tapetum of the Potamogeton species studied here was classified as plasmodial due to the loss of the cell walls and fusion of the protoplasts, in accordance with Furness and Rudall (2001). This finding is also consistent with the tapetum type of Alismatales, except for Tofieldia (Toelfidiaceae) (Furness and Rudall, 1998). The tapetal cells of the species studied were binucleate, an unusual
characteristic in this genus, reported only for P. natans (Haynes and Holm-Nielsen, 2003). Stevens (2009) reported uninucleate tapetum cells as a synapomorphy of Alismatales. The curvature of the Potamogeton ovule has been the subject of differing interpretations. According to Davis (1966) and Igersheim et al. (2001), the ovule in Potamogeton is orthotropous. An alternative description, based on the ovule development of P. natans, was proposed by Takaso and Bouman (1984). These authors classified the ovule in this species as ortho-campylotropous, since the primordium was oriented almost horizontally, becoming campylotropous only after megasporogenesis. The ovule of Potamogeton has also been defined as anatropous (Posluszny and Sattler, 1974; Posluszny, 1981). Our observations do not support the interpretation of orthotropous curvature, since the Potamogeton ovule is not completely straight and because its chalaza, nucellus and micropyle are not aligned. In fact, the curvature of the ovule and the inversion of nucellus reported here are not caused solely by intercalary growth of the funiculus. The mature megagametophyte is curved and the integuments are similarly developed on both the abaxial
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and adaxial sides. The ovule was interpreted as campylotropous in the present investigation, since the megagametophyte is curved at maturity (Corner, 1976). In the three species studied, curvature begins before megasporogenesis with increased development in the funiculus and chalaza regions. The proliferating tissue in the proximal region enhances the campylotropous curvature in Potamogeton. This interpretation is in accordance with Bouquet and Bersier (1960) and corroborates the observations of Takaso and Bouman (1984) of the P. natans ovule. Dahlgren et al. (1985) identified only two characters relating to the micropylar region in monocotyledons: (1) the presence or absence of the parietal cells or (2) the presence or absence of a multiseriate nucellar cap. Both cases in Potamogeton species occur: the parietal cell forms parietal layers and the nucellar epidermis forms a nucellar cap. Although Wiegand (1900) has described only the occurrence of parietal layers in P. foliosus, Takaso and Bouman (1984) found both phenomena in P. natans. A nucellar cap has also been observed in Araceae (Grayum, 1991) and Acorus (Acoraceae) (Rudall and Furness, 1997), as well in some magnoliids (Tobe et al., 1993; Tobe and Sampson, 2000; Kimoto and Tobe, 2001). Based on these data, Tobe (2008) suggested that a twocell-layered nucellar cap is a possible plesiomorphy in monocots and could represent a morphological link between monocots and magnoliids. During megasporogenesis, the micropylar dyad fails to complete meiosis II, resulting in a triad of megaspores. This feature is at odds with the occurrence of T-shaped tetrads reported by Johri et al. (1992). The former phenomenon was observed in P. natans by Takaso and Bouman (1984) and in Tofieldia glutinosa (Tofieldiaceae) by Holloway and Friedman (2008). Tofieldiaceae has recently been positioned within the Alismatales by molecular phylogenetic analysis (APG II, 2003; Soltis et al., 2007; APG III, 2009). Although this aspect of megasporogenesis may seem to be a rare phenomenon, it is more widespread than might be assumed, having been reported in species of Anacardiaceae (Grundwag, 1976), Austrobaileyaceae (Tobe et al., 2007), Brassicaceae (Spooner, 1984), Leguminosae (Seshavatharam and Subba Rao, 1982; Rodrígues˜ et al., 2006), Nymphaeaceae (Orban and Bouharmont, 1998; Riano Friedman and Williams, 2003), and Orchidaceae (Abe, 1976; Law and Yeung, 1989). This study provides new observations about the embryology of Potamogeton, such as monocotyledonous-type anther wall development, pectic–cellulosic thickenings in the endothecium, microspore tetrads (most commonly tetragonal and decussate), campylotropous ovule, and triads of megaspores resulting from meiosis II failure in the micropylar dyad cell. These data were consistent and conserved across the three species studied, despite their positioning in different clades by molecular phylogenetic analysis (Lindqvist et al., 2006). The following observations differ from those compiled by Johri et al. (1992) for Potamogetonaceae: endothecium with pectic–cellulosic thickenings, predominantly tetragonal and decussate microspore tetrads, inaperturate pollen grains, with reticulate exine and dispersed with three cells, and campylotropous ovule. Such differences are probably due to the modifications in the circumscription of the family following molecular phylogenetic analyses in recent years.
Acknowledgements We thank the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq 47248/2004-0) and the Coordenac¸ão de Aperfeic¸oamento de Pessoal de Nível Superior (CAPES) (MSc grant to E.L.P.N.) for their financial support. We acknowledge Instituto Ecoplan (PR, Brazil) and Indústrias Pizzatto (PR, Brazil) for logistic support.
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Contribution to the embryology of Potamogeton L. (Alismatales: Potamogetonaceae) Elaine Lopes Pereira Nunes ∗ , Mariana Cortes de Lima, Alessandra Ike Coan 1 , Maria Cecília de Chiara Moc¸o 2 Departamento de Botânica, Setor de Ciências Biológicas, Universidade Federal do Paraná, C. Postal 19031, CEP 81531-980, Curitiba, Paraná, Brazil
a r t i c l e
i n f o
Article history: Received 3 November 2009 Received in revised form 24 February 2010 Accepted 4 March 2010 Available online 10 March 2010 Keywords: Anther wall Megagametogenesis Megasporogenesis Microgametogenesis Microsporogenesis Monocotyledon
a b s t r a c t Potamogetonaceae currently comprises four genera, including Potamogeton, a near cosmopolitan genus of about 100 species. The historical circumscription of this family is controversial, resulting in fragmentary information on the morphology and anatomy of its genera. The aim of the present study was to investigate the anther and ovule development in three species of Potamogeton in order to clarify the embryological features of this genus. The majority of embryological characters were constant among the three species studied and are in accordance with previous reports. In addition, some new and complementary characteristics of Potamogeton embryology are highlighted, including: monocotyledonous-type of anther wall development and carpels bearing a single ovule that develops a Polygonum-type of megagametophyte. A fail in the micropylar dyad to divide is confirmed as the cause of megaspore triads. The developmental sequence of the ovule confirms its campylotropous curvature. © 2010 Elsevier B.V. All rights reserved.
1. Introduction Potamogetonaceae is a subcosmopolitan and aquatic family (Haynes and Holm-Nielsen, 2003) currently placed within the early-divergent order Alismatales (APG III, 2009). This family has previously been placed in the Helobiae (Engler, 1904), Potamogetonales (Takhtajan, 1997), and Najadales (Cronquist, 1981), based mainly on morphological characters. The circumscription of genera within Potamogetonaceae has also varied among different authors over time. The family has been combined with members of Cymodoceaceae, Najadaceae, Zannichelliaceae and Zosteraceae (Haynes and Holm-Nielsen, 2003). According to Haynes and Holm-Nielsen (2003), the genera included in Potamogetonaceae are: Groenlandia J. Gay, Potamogeton L. and Stuckenia Börner. However, APG II (2003) has merged Zannichelliaceae in Potamogetonaceae, with the addition of three more genera: Althenia Petit, Lepilaena J.Drum. ex Harv and Zannichellia L. More recently, Lindqvist et al. (2006), using non-coding nuclear and plastid DNA data to determine Potamoge-
∗ Corresponding author. Tel.: +55 41 3361 1620; fax: +55 41 3266 2042. E-mail address: [email protected] (E.L.P. Nunes). 1 Present address: Departamento de Botânica, Instituto de Biociências, Universidade Estadual Paulista “Júlio de Mesquita Filho”, C. Postal 199, CEP 13506-900, Rio Claro, São Paulo, Brazil. 2 Present address: Departamento de Botânica, Instituto de Biociências, Universidade Federal do Rio Grande do Sul, Av. Bento Gonc¸alves, 9500, CEP 91509-900, Porto Alegre, Rio Grande do Sul, Brazil. 0304-3770/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.aquabot.2010.03.006
tonaceae relationships, concluded that Potamogeton, Groenlandia and Stuckenia, plus Zannichellia L., form a strongly supported monophyletic group. Potamogeton is the largest genus of Potamogetonaceae, with about 100 species (Haynes and Holm-Nielsen, 2003). It is near cosmopolitan, herbaceous, and dispersed by seeds, turions or rhizomes (Haynes and Holm-Nielsen, 2003). Because of its high phenotypic plasticity, hybridization, polyploidy and aneuploidy (Hollingsworth et al., 1998; Kaplan, 2002; Wang et al., 2007), Potamogeton has been historically divided into many infrageneric categories (e.g., Ascherson and Graebner, 1907; Hagström, 1916; Fernald, 1932; Ogden, 1943). However, no such categories were adopted in more recent taxonomic work on Potamogeton (Wiegleb and Kaplan, 1998), which only included notes on systematic affinities. Similarly, the systematic treatment of Potamogetonaceae for the Flora of North America (Haynes and Hellquist, 2000) and for the Flora Neotropica (Haynes and Holm-Nielsen, 2003) did not include an infrageneric classification. A review of the literature shows that the embryological features of Potamogetonaceae are unstable, mainly because of the aforementioned differences in circumscription. The embryological characters compiled by Johri et al. (1992) included genera currently assigned to other families, such as Syringodium Kütz. (Cymodoceaceae), Phyllospadix Hook. and Zostera L. (Zosteraceae). The correct correspondence of each embryological character to each of the above genera is therefore not clear, even for Potamogeton.
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A summary of embryological characters known for Potamogeton is as follows: endothecium with fibrous thickenings (Gupta, 1934; Johri et al., 1992); microsporogenesis of the successive type (Furness and Rudall, 1999 based on Stenar, 1925 and Schnarf, 1931); ovule primordium with three zones (Takaso and Bouman, 1984); ovule ortho-campylotropous (Takaso and Bouman, 1984), orthotropous (Davis, 1966; Igersheim et al., 2001), or anatropous (Posluszny and Sattler, 1974; Posluszny, 1981), bitegmic and crassinucelate (Takaso and Bouman, 1984); linear or T-shaped arrangements of megaspore tetrads (Johri et al., 1992) and megagametophyte of the Polygonum-type (Takaso and Bouman, 1984). According to Haynes and Holm-Nielsen (2003), the type of anther wall development is unknown. Because of the confusion caused by successive circumscription changes in Potamogetonaceae, the aim of the present study was to investigate the early development of the anther and carpel, focusing on micro- and mega-sporogenesis and gametogenesis, of three species of Potamogeton in order to gain a better understanding of the embryology of the genus.
2. Materials and methods Inflorescences at successive developmental stages of three species of Potamogeton were collected in lakes and rivers of Brazil. Vouchers were deposited in the Herbário do Departamento de Botânica, Universidade Federal do Paraná, Curitiba, Paraná, Brazil (UPCB): Potamogeton illinoensis Morong (UPCB 60501), Potamogeton polygonus Cham. & Schltdl (UPCB 61311), and Potamogeton pusillus L. (UPCB 60500). For light microscope (LM) examination, the material was fixed in 1% glutaraldehyde and 4% formaldehyde in 0.1 M phosphate buffer, pH 7.2 (McDowell and Trump, 1976). Samples were rinsed in phosphate buffer, dehydrated through a graded ethanol series and embedded in Leica historesin. Sections (2–5 m) were cut on a Leica RM2145 microtome using glass knives, stained with toluidine blue (O’Brien et al., 1965) and mounted in Permount. Photomicrographs were taken using a Zeiss Axiolab with a digital camera. The following stains were performed on resin-embedded semi-thin sections: iodine potassium iodide solution for starch (Johansen, 1940), Coomassie Brilliant Blue for protein (Southworth, 1973), Ruthenium Red for pectin (Jensen, 1962), Calcofluor White MR2 for cellulose (Feder and O’Brien, 1968) and Aniline Blue for callose (modified from Arens, 1949). Callose and cellulose epifluorescence was observed using a Zeiss Axiophot microscope with a DAPI filter set. Fixed anthers were stained for lipids with Sudan III (O’Brien and McCully, 1981) and for lignin with acid phloroglucin (Sass, 1951). Fixed mature anthers were macerated for 4 h at 60 ◦ C in acetic acid and hydrogen peroxide (1H2 O2 (30%):4 distillate water:5 glacial acetic acid; modified from Ruzin, 1999) and mounted in glycerin jelly to allow observation of the three-dimensional structure of endothecium.
3. Results 3.1. Anther development The flowers of Potamogeton are composed of four tepals, four stamens and four free carpels. In longitudinal sections of the flower, each anther primordium arises as a small group of undifferentiated cells above each tepal primordium (Fig. 1A and B). Protodermal cells give rise to the epidermis through anticlinal divisions (Fig. 1C and D). Archesporial cells are produced by the ground meristem (Fig. 1B and C).
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The development of the anther wall is initiated by the differentiation of the archesporial cells into sporogenous and primary parietal cells (Fig. 1D – arrow). The primary parietal cells undergo periclinal division to produce two secondary parietal layers (Fig. 1E – arrow). The outer layer develops directly into the endothecium, and the inner layer undergoes periclinal divisions to originate two to three ephemerous middle layers and the tapetum (Fig. 1F – arrow and G). The sporogenous cells differentiate into microsporocytes soon after some mitotic divisions (Fig. 1E–G). During meiosis in microsporocytes, the middle layers are compressed against the endothecium and are no longer apparent in the mature anther (Fig. 1H and I). The tapetum is initially a singlelayered tissue with uninucleate cells (Fig. 1F), except in P. polygonus, whose cells become binucleate slightly before they detach one from another (Fig. 1H and I). In P. illinoensis, some cells of the tapetum divide further, giving rise to up to four layers (not shown). The tapetum is one of the plasmodial type, since its cells lose their walls and migrate into the anther locule (Figs. 1H and I and 2A and B). A periplasmodium was observed until the free microspore stage, and disappeared before the onset of microgametogenesis (Fig. 2B). Microsporocytes are rounded (Fig. 1G) and start meiosis concurrent with the detachment of tapetum cells (Fig. 1H and I). The microsporocytes undergo meiosis I as usual (Fig. 1G–I), followed by cytokinesis and dyad formation (Fig. 1I). After meiosis II, a wide range of tetrad shapes was observed, the most common being the tetragonal and decussate (Fig. 2A and B; detailed observations on this issue are discussed by Nunes et al., 2009). Microgametogenesis begins with the formation of a large vacuole that pushes the nucleus towards the cell wall (Fig. 2C) followed by unequal mitotic division of the microspore, resulting in a small generative cell and a large vegetative cell (Fig. 2D). Before shedding, the generative nucleus divides to form two small sperm cells (Fig. 2E). The cytoplasm of the vegetative cell contains numerous starch grains (Fig. 2E). The mature pollen grain has no apertures. The exine is thin and the intine is pectic–cellulosic in nature. The endothecium has large polyhedral cells and Ushaped pectic–cellulosic thickenings by the time of dehiscence (Fig. 2F). Each microsporangium pair coalesces before pollen shedding (Fig. 2F) and dehiscence is longitudinal. 3.2. Ovule development The gynoecium consists of four carpels in a single whorl (Fig. 1A). A single three-zonate ovule primordium is initiated in the ventral wall of each carpel (Fig. 3A). The ovule develops almost horizontally, filling the locule (Fig. 3B and C). Two integuments with two rows of cells each are formed, eventually expanding to three rows, especially next to the chalaza (Fig. 3B–D). As the ovule develops, it gradually bends down to the carpelary base (Fig. 3C–F). This campylotropous curvature is initially brought about by two processes: increased growth of the adjacent region between the chalaza and the integuments, and growth of the carpel, creating space to accommodate the ovule (Fig. 3C–F – marked area). The micropyle is formed by the inner integument alone (Fig. 3G). At the time the megagametophyte is mature, the median region of the ventral wall of the carpel proliferates near the funiculus (Fig. 3H – arrow). This proliferating tissue puts pressure on the funicular region and accentuates the curvature of the ovule. The nucellus divides periclinally, giving rise to four or five parietal layers. The mature ovule is campylotropous, bitegmic and crassinucelate (Fig. 3F–H). The megasporocyte differentiates after the inception of integuments (Fig. 3C). Meiosis I produces a dyad of cells; the micropylar cell is smaller than the chalazal (not shown). Meiosis II proceeds normally in the chalazal cell, producing two megaspores (Fig. 4A). In the micropylar cell, meiosis II is disrupted (Fig. 4A – arrow). This cell soon degenerates, followed by the adjacent megaspore (Fig. 4B
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Fig. 1. Early stages of anther development in P. polygonus (A–F and H–I) and P. illinoensis (G). (A) Longitudinal section (LS) of floral bud showing whorls inception. (B) LS of floral bud with anther primordium in detail. (C) Transversal section (TS) of anther, showing tissues differentiation. (D) Anther primordium showing primary parietal layer (arrow) and sporogenous tissue differentiated, in TS. (E) Periclinal division of the primary parietal layer (arrow), in TS. (F) Anther after a periclinal division of the inner secondary parietal layer (arrow), in TS. (G) Pre-meiotic anther with parietal layers and microsporocytes differentiated, in TS. (H) Anaphase I of meiosis and detachment of binucleate tapetal cells, in TS. (I) Microspore dyads in telophase I. C: carpel; e: epidermis; en: endothecium; MI: microsporocytes; ml: middle layers; pc: procambium; pd: protodermis; S: stamen; sc: sporogenous cells; T: tepal; t: tapetum. Scale bars = 10 m in (A); 20 m in the remaining images.
– arrows). No fully developed megaspore tetrads were observed in any species. Callose was detected only in the transversal walls of the triad. The chalazal-most megaspore is functional (Fig. 4B). Megagametogenesis is brought about by three cycles of mitotic division, resulting in a Polygonum-type megagametophyte (Fig. 4C–E and G–I). When mature, the megagametophyte consists of seven cells: three ephemeral antipodal cells at the chalazal pole (Fig. 4H); the central cell with two polar nuclei that fuse prior to fertilization (Fig. 4H); a single egg cell (Fig. 4G) and two synergids (Fig. 4I) at the micropylar pole. During megasporogenesis and megagametogenesis, the apical epidermal cells and possibly some hypodermal cells of the nucellus divide periclinally to form a nucellar cap (Fig. 4A–F). Neither a hypostase nor an obturator is formed during ovule development.
4. Discussion Anther wall development of Potamogeton is described in detail for the first time in this report. According to the features observed in P. illinoensis, P. polygonus and P. pusillus, anther wall development is of the monocotyledonous-type, in which the inner secondary parietal layer divides to give rise to the middle layers and the tapetum, in accordance with Davis (1966). In the present study, the development of U-shaped pectic–cellulosic thickenings in the endothecial cells was observed at the free microspore stage. Although the presence of thickenings is in agreement with the available data for Potamogeton (Johri et al., 1992; Haynes and Holm-Nielsen, 2003), their nature has remained unclear until now. The type of thickening observed here
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Fig. 2. Late microsporogenesis and microgametogenesis in Potamogeton polygonus (A and C–F) and P. pusillus (B). (A) Microspore tetrads in telophase II and binucleate tapetum cell migrating to the anther locule. (B) Microspore tetrads enclosed by a periplasmodium, in TS. (C) Microgametogenesis onset. (D) Young bicellular pollen grain. (E) Tricellular pollen grain in a mature anther. (F) Anther thecae before dehiscence, in TS. en: endothecium; gc: generative cell; sp: sperm cells; vc: vegetative cell; t: tapetum. Scale bars = 30 m in (J); 10 m in the remaining images.
Fig. 3. Ovule development in Potamogeton polygonus (A, B and D–H) and P. iilinoensis (C). (A) Carpel primordium, in longitudinal section (LS), showing an ovule primordium with three zones. (B) Transversal section (TS) of a carpel with an ovule primordium after the inception of the integuments. (C) Ovule primordium before megasporocyte meiosis in the beginning of its curvature; the development is greater at the interface between chalaza and integuments, in LS. (D–F) Successive stages of ovule curvature due to the greater development of the interface (marked area) between the chalaza and the integuments, in LS. (G) Detail of the micropyle (arrow) of a mature ovule, in LS. (H) LS of ovary showing mature campylotropous ovule, note that the ventral wall of the carpel, adjacent to the the funiculus (arrow), proliferates. ii: inner integument; dw: dorsal wall; n: nucellus; ov: ovule; oi: outer integument; vb: vascular bundle; vw: ventral wall. Scale bars = 50 m in (F) and (H); 20 m in the remaining images.
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Fig. 4. Megasporogenesis and megagametogenesis in Potamogeton polygonus (A–I), ovule in longitudinal sections. (A) Chalazal megaspore dyad and degenerated micropylar dyad (arrow). (B) Functional megaspore at the chalazal pole and degenerated micropylar dyad and adjacent megaspore (arrows). (C) Functional megaspore with cell content at its micropylar region. (D) Immature binucleate megagametophyte (only one nucleus is visible). (E) Four-nucleate megagametophyte (only the two chalazal nuclei are visible). (F) Detail of the micropyle showing the nucellar cap (arrow). (G) Megagametophyte detail showing the egg cell and a synergid partially visible. (H) Megagametophyte detail showing the antipodal cells and the central cell. (I) Polygonum-type megagametophyte. an: antipodals; cc: central cell; ec: egg cell; ii: inner integument; n: nucellus; oi: outer integument; sy: synergid. Scale bars = 50 m in (D) and (E); 10 m in (G) and (H); 20 m in the remaining images.
contrasts with the girdle-type reported by Johri et al. (1992) and Haynes and Holm-Nielsen (2003). In this study, all histochemical staining tests indicated that the endothecial thickenings are definitely composed of pectins and cellulose, but not guaiacyl lignins. However, a negative result of the phloroglucinol–HCl test does not rule out the presence of lignin. Generally, lignins are classified as guaiacyl, guaiacyl–syringyl, or guaiacyl–syringyl–phydroxyphenyl lignins, according to whether they are from gymnosperms, wood angiosperms, or grasses, respectively. A positive result of the phloroglucinol–HCl indicates the presence of coniferaldehyde end-groups in guaiacyl lignins only. However, Lewis and Yamamoto (1990) emphasized that there are exceptions, and that no single analytical method provides conclusive evidence that lignin is present. The tapetum of the Potamogeton species studied here was classified as plasmodial due to the loss of the cell walls and fusion of the protoplasts, in accordance with Furness and Rudall (2001). This finding is also consistent with the tapetum type of Alismatales, except for Tofieldia (Toelfidiaceae) (Furness and Rudall, 1998). The tapetal cells of the species studied were binucleate, an unusual
characteristic in this genus, reported only for P. natans (Haynes and Holm-Nielsen, 2003). Stevens (2009) reported uninucleate tapetum cells as a synapomorphy of Alismatales. The curvature of the Potamogeton ovule has been the subject of differing interpretations. According to Davis (1966) and Igersheim et al. (2001), the ovule in Potamogeton is orthotropous. An alternative description, based on the ovule development of P. natans, was proposed by Takaso and Bouman (1984). These authors classified the ovule in this species as ortho-campylotropous, since the primordium was oriented almost horizontally, becoming campylotropous only after megasporogenesis. The ovule of Potamogeton has also been defined as anatropous (Posluszny and Sattler, 1974; Posluszny, 1981). Our observations do not support the interpretation of orthotropous curvature, since the Potamogeton ovule is not completely straight and because its chalaza, nucellus and micropyle are not aligned. In fact, the curvature of the ovule and the inversion of nucellus reported here are not caused solely by intercalary growth of the funiculus. The mature megagametophyte is curved and the integuments are similarly developed on both the abaxial
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and adaxial sides. The ovule was interpreted as campylotropous in the present investigation, since the megagametophyte is curved at maturity (Corner, 1976). In the three species studied, curvature begins before megasporogenesis with increased development in the funiculus and chalaza regions. The proliferating tissue in the proximal region enhances the campylotropous curvature in Potamogeton. This interpretation is in accordance with Bouquet and Bersier (1960) and corroborates the observations of Takaso and Bouman (1984) of the P. natans ovule. Dahlgren et al. (1985) identified only two characters relating to the micropylar region in monocotyledons: (1) the presence or absence of the parietal cells or (2) the presence or absence of a multiseriate nucellar cap. Both cases in Potamogeton species occur: the parietal cell forms parietal layers and the nucellar epidermis forms a nucellar cap. Although Wiegand (1900) has described only the occurrence of parietal layers in P. foliosus, Takaso and Bouman (1984) found both phenomena in P. natans. A nucellar cap has also been observed in Araceae (Grayum, 1991) and Acorus (Acoraceae) (Rudall and Furness, 1997), as well in some magnoliids (Tobe et al., 1993; Tobe and Sampson, 2000; Kimoto and Tobe, 2001). Based on these data, Tobe (2008) suggested that a twocell-layered nucellar cap is a possible plesiomorphy in monocots and could represent a morphological link between monocots and magnoliids. During megasporogenesis, the micropylar dyad fails to complete meiosis II, resulting in a triad of megaspores. This feature is at odds with the occurrence of T-shaped tetrads reported by Johri et al. (1992). The former phenomenon was observed in P. natans by Takaso and Bouman (1984) and in Tofieldia glutinosa (Tofieldiaceae) by Holloway and Friedman (2008). Tofieldiaceae has recently been positioned within the Alismatales by molecular phylogenetic analysis (APG II, 2003; Soltis et al., 2007; APG III, 2009). Although this aspect of megasporogenesis may seem to be a rare phenomenon, it is more widespread than might be assumed, having been reported in species of Anacardiaceae (Grundwag, 1976), Austrobaileyaceae (Tobe et al., 2007), Brassicaceae (Spooner, 1984), Leguminosae (Seshavatharam and Subba Rao, 1982; Rodrígues˜ et al., 2006), Nymphaeaceae (Orban and Bouharmont, 1998; Riano Friedman and Williams, 2003), and Orchidaceae (Abe, 1976; Law and Yeung, 1989). This study provides new observations about the embryology of Potamogeton, such as monocotyledonous-type anther wall development, pectic–cellulosic thickenings in the endothecium, microspore tetrads (most commonly tetragonal and decussate), campylotropous ovule, and triads of megaspores resulting from meiosis II failure in the micropylar dyad cell. These data were consistent and conserved across the three species studied, despite their positioning in different clades by molecular phylogenetic analysis (Lindqvist et al., 2006). The following observations differ from those compiled by Johri et al. (1992) for Potamogetonaceae: endothecium with pectic–cellulosic thickenings, predominantly tetragonal and decussate microspore tetrads, inaperturate pollen grains, with reticulate exine and dispersed with three cells, and campylotropous ovule. Such differences are probably due to the modifications in the circumscription of the family following molecular phylogenetic analyses in recent years.
Acknowledgements We thank the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq 47248/2004-0) and the Coordenac¸ão de Aperfeic¸oamento de Pessoal de Nível Superior (CAPES) (MSc grant to E.L.P.N.) for their financial support. We acknowledge Instituto Ecoplan (PR, Brazil) and Indústrias Pizzatto (PR, Brazil) for logistic support.
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