Sporogenesis in bryophytes: patterns and diversity in meiosis.
The sphagnum mosses (Sphagnopsida) are an early divergent and distinct group. They are hydrophytes of worldwide distribution, but more abundant in boreal regions of the world where they are a major player in the succession of bogs. Sphagnum is of ecological importance in habitat formation and as a sensitive indicator of pollution. They produce a surprising amount of biomass and in wet northern habitats are known as peat moss. Incomplete decomposition under limited oxygen conditions leads to build up of the organic matter known as peat. Peat lands occupy about 4% of the world's surface (Turner, 1993). Peat is of considerable economic importance and with diverse applications such as a low-grade source of fuel for heating, in distilling of Scotch whiskey, as a horticultural growing medium and soil supplement, and as a source of waxes and resins. Again the history of exploitation and xerification in general would suggest that these ancient, biologically fascinating and economically valuable organisms are in peril. A recent volume of Advances in Bryology (Miller, 1993) is dedicated to the biology of these remarkable organisms.
Undeniably monophyletic, they are a large taxonomically complex group with obscure relationships to other mosses. Morphological evidence emphasizes the isolation of the group within the Bryophytes. More classical (Crum & Anderson, 1981) as well as modem classifications (Goffinet & Shaw, 2009) treat the sphagnum mosses as a monotypic class, the Sphagnopsida.
Not surprisingly, spore and spore dispersal mechanisms in sphagnum are profoundly different from other mosses (e.g. Duckett et al., 2009). Sphagnum bas a peculiar manner of spore release by explosive de-operculation of the capsule. The spores of Sphagnum are uniquely multilayered and omate (Brown et al., 1982a, 1982b). Spores are trilete, with strongly differentiated proximal and distal surfaces. The two hemispheres are separated by a distinct rim or cingulum at the equator. The proximal face has a conspicuous trilaesurate aperture and the distal hemisphere is ornamented with a radiating pattern of ridges. The trilete proximal aperture, a feature shared with Takakia (Renzaglia et al., 1997) and Oedopodium (Shimamura & Deguchi, 2008), is not characteristic of the peristomate mosses where spores are typically hilate. The functional trilete aperture of Sphagnum is more characteristic of free-sporing vascular plants than of mosses.
Meiosis in Sphagnum provides a dramatic example of the involvement of the QMS in establishment of division polarity and spindle ontogeny. The meiotic spindle remains quadripolar, its poles anchored at the plastids positioned in tetrahedral arrangement. This remarkable structure delivers two groups of chromosomes to the polar cleavage furrows. The telophase I nuclei remain elongated and stretch between the pairs of poles on either side of the polar cleavage furrow. This differs from other bryophytes with monoplastidic meiosis where the four poles of the QMS detach from the plastids and converge in pairs toward the polar cleavage furrows.
The early prophase sporocyte is polarized; one hemisphere contains the acentric nucleus while two sausage shaped plastids are positioned in the other (Figs. 58a and 59a). The two plastids are interconnected by microtubules that emanate from the plastid surfaces (Fig. 58a). The two plastids begin to rotate so that their tips are in tetrahedral arrangement while the nucleus is still acentric (Fig. 59a). By midprophase the nucleus is centered in the sporocyte, the plastids have divided and the four resulting plastids are positioned at the tetrad poles (Fig. 58b-d and 59b) and interconnected by the QMS (Fig. 58b, c). Sphagnum provides an outstanding example of the unusual quadripolarity of the metaphase I spindle that can be traced directly to the plastid based QMS of prophase. The quadripolar prometaphase spindle (Fig. 58e, f) develops as microtubules from the four plastid poles, one shown in Fig. 59c, become more numerous and ordered into fibers.
In metaphase I (Figs. 58g, h, 59d) kinetochore bundles extend between the chromosomes to broad polar regions arching around polar cleavage furrows. The spindle remains quadripolar; the two half spindles are shaped like a pair of axe heads at right angles to each other with a plastid at each of the corners (Fig. 58g, h). In a strictly equatorial view, the spindle appears as a triangle (Fig. 59d). The spindle poles come to a sharp edge (Figs. 58g and 6la). Elements of ER that parallel the microtubules tend to be more associated with the polar regions (Fig. 6la). Such ER may be transported to the poles and contribute to reformation of the nuclear envelopes. In anaphase I (Fig. 58i, j), the kinetochore fibers shorten. The spindle remains suspended from the four plastids at the comers and chromosomes are distributed along the broad arcs of the polar regions in telophase I. Interzonal micrombules, which proliferate from proximal surfaces of reforming nuclei and plastids, forma conspicuous phragmoplast (Fig. 58k, 1), which dissipates without cell plate deposition. Sporogenesis differs from bryopsid mosses in that there is no conspicuous organelle band separating the dyad domains, although it is possible that mitochondria are preferentially located in the equatorial region (Fig. 61b). The plastids remain at the tetrad poles and numerous oil bodies are dispersed throughout the cytoplasm.
The second meiotic division occurs simultaneously in the undivided cytoplasm with spindle orientation resulting in the placement of a nucleus adjacent to each of the four plastids at the tetrad poles (Figs. 60a h and 61b-d). The elongated dyad nuclei are suspended between plastids at approximately right angles with tips in tetrahedral arrangement (Fig. 60a, b). Microtubules emanate from the four plastid MTOCs to re-establish a tetrahedral QMS (Fig. 60a). Some of the opposing microtubules envelope the dyad nuclei resulting in prophase spindles (Fig. 60a) and others interconnect the plastids in bipolar phragmoplast-like arrays (Fig. 60a). The metaphase II spindles (Fig. 60c, d and 61b) terminate at the plastids. The influence of quadripolarity on spindle morphology can be seen by comparing metaphase I and II spindles in a fortuitous arrangement of two sporocytes (Fig. 62). Anaphase II spindles (Fig. 60e, t) appear completely orthodox with pointed poles. Cytokinesis is simultaneous. Primary phragmoplasts are initiated in the interzones of spindles and additional microtubules emanating from non-sister nuclei/plastids give rise to secondary phragmoplasts that interconnect the nuclei and define the division planes (Fig. 60g, h). Numerous vesicles accumulate at the interfaces of the bipolar arrays and contribute to the intersporal cell plates (Fig. 61c, d); the developing intersporal septa join precisely with the prophasic ingrowths of the sporocyte wall.
Meiosis in Entodon seductrix
Sporocytes of the bryopsid mosses produce isolating mucopolysaccharide walls within cell walls of the archesporium. The cellulosic walls subsequently lyse and the sporocytes are released into a common chamber, the so-called spore sac (Fig. 63). As in Sphagnum, the number of plastids is reduced to one in each sporocyte.
The single plastid (Fig. 64a) divides once resulting in sporocytes with two plastids while the nucleus is still acentric (Figs. 64a and 65a). An extensive plastid based microtubule system (Figs. 64a-e and 65b) is associated with each of the plastids. This microtubule system resembles the AMS described in meiosis of hornworts in that it is oriented parallel to the long axis of the plastids (Figs. 64a and 65b). The nature of the AMS is especially obvious in the late dividing plastids of the moss Funaria. Tips of the two plastids are at the vertices of a tetrahedron and serve as the MTOCs that nucleate microtubules of the QMS, but the plastids do not actually complete division until metaphase I (Busby & Gunning, 1988a, b). In E. seductrix the two plastids divide again and the four plastids are positioned in tetrahedral arrangement where they are poles of the QMS during mid-late prophase (Figs. 64c-e and 65c). The QMS interconnects the plastids and encages the nucleus in four interacting cones of microtubules (Fig. 64c-e). The nucleus itself becomes tetrahedrally lobed as it is drawn to the microtubule foci (Figs. 64d and 65c). Increased vesicular activity is apparent at the interfaces of the opposing arrays (Fig. 65c, d) where additional sporocyte wall material is deposited, thus deepening the infurrows and contributing to lobing of the sporocyte into the four future spore domains. These rings of vesicles coincide with the interface of opposing cones of microtubules to define polar and equatorial division planes at the boundaries of the spore domains.
The meiotic spindle is quadripolar in origin and develops from convergence of pairs of poles of the QMS in prometaphase I (Fig. 64f-h). Compare the QMSs shown in adjacent sporocytes (Fig. 64f) to the prometaphase spindle (Fig. 64g, h). As seen in equatorial view (Figs. 64g, h and 66a), each pair of poles is adjacent to a polar cleavage furrow. In metaphase I, the spindle poles arch around the cleavage furrows (Fig. 64i, j). All signs of direct microtubule attachment to plastids disappear and the microtubule foci move toward the cleavage furrows. In most cases, there is no evidence of a discrete structure associated with the poles, but in the bryopsid moss Rhynchostegium serrulatum, spindle microtubules appear to be focused at electron dense clusters (Brown and Lemmon, 1982a) that are thought to constitute the MTOCs. In anaphase I (Fig. 64k, 1) the spindle poles at cleavage furrows become more consolidated.
Telophase I nuclei are delivered to opposite polar cleavage furrows (Fig. 66b). Immediately a distinctive phragmoplast develops (Fig. 67a), however an intersporal wall does not develop. Instead another distinctive and unusual component of organellar mitosis occurs, the development of the organelle band (OB) (Fig. 66c). The OB is a distinctive structure that occurs as a component of meiosis in all major groups of land plants. In mosses as in other plants with monoplastidic meiosis, the plastids remain locked in position at tetrad poles where
they function as poles for the second division, which occurs in dyad domains on either side of the OB. In telophase I the larger organelles associate with the vesicles of the equatorial infurrows forming a collar around the spindle (Fig. 66b). They invade the interzonal region and form a plate-like aggregation of vesicles, ER fragments, mitochondria, and other organelles such as oil bodies (Fig. 66c). While vesicles and ER fragments are thought to be transported to the forming cell plate by the phragmoplast, it is not at all clear how large organelles such as oil bodies and mitochondria move into the equatorial plane, especially as the movement appears to be at fight angles to the orientation of the phragmoplast microtubules. The OB is thought to function in separating the second division spindles and/or assuring equal apportionment of organelles to the four spore domains.
In prophase (Fig. 67b, c), each nucleus is elongated in the spindle axis between a pair of plastids. The nuclei are shrouded by microtubules emanating from the plastid MTOCs. This results in a pair of metaphase II spindles with poles adjacent to the plastids in tetrahedral arrangement (Fig. 67d). Flattening during preparation can produce a pair of second division spindles in a cross-like arrangement (Fig. 67e, f). Simultaneous mitosis results in each spore domain receiving a nucleus adjacent to the single plastid (Fig. 67g, h). Phragmoplasts develop between pairs of telophase II nuclei (Fig. 67g, h). As the primary phragmoplasts expand additional microtubules from the plastid MTOCs interact and initiate secondary phragmoplasts (Fig. 67i, j). Deposition of intersporal walls begins in the primary phragmoplasts while the first division site is still occupied by the OB (Fig. 66d). The intersporal walls fuse with the peripheral wall ingrowths formed during prophase I and the components of the OB are equally distributed to the tetrad members. In young spores of the tetrad, microtubules radiate from a discrete MTOC (Fig. 67k, 1). Nothing is known of the mechanisms of its relocation, but following exine initiation the MTOC migrates to the proximal surface where it is associated with the localized deposition of the hilate aperture.
Meiosis in Trematodon longicollis
Localization of [gamma]-tubulin throughout the process of monoplastidic meiosis in T. longicollis augments information from light and electron microscopy. Triple staining for microtubules, [gamma]-tubulin and nuclei provides the first complete account of the migrations and form changes that [gamma]-tubulin regularly undergoes during monoplastidic meiosis in mosses. The dynamics of the plastids can be see in both the [gamma]-tubulin channel where they are invested with [gamma]-tubulin and the nuclear channel where they show a lighter staining than the nucleus. The fundamental role of the QMS establishes ail aspects of division quadripolarity; it determines the future cytokinetic planes and spindle orientation for both meiosis I and II. Although the plastid MTOCs control quadripolarity and generate the spindles, the behavior of [gamma]-tubulin includes a series of precisely programmed migrations away from the plastids during meiosis I and back to the plastids in meiosis II, followed by a condensation and dramatic migration to the site of aperture development in the four spores following cytokinesis.
The early unlobed sporocytes contain four plastids (Fig. 68a-c) resulting from two divisions of the single plastid. The plastids, not yet in tetrahedral position, are MTOCs as evidenced by the presence of [gamma]-tubulin (Fig. 68b) and microtubules emanating from them (Fig. 68a). The bipolar arrays between daughter plastids are slightly more prominent (Fig. 68a) than the remaining microtubules. At a slightly later stage in early prophase, the plastids are positioned at the tetrad poles and both cytoplasm and nucleus are tetrahedrally lobed (Fig. 68d-f). Microtubules emanating from the plastid MTOCs organize the complex QMS (Fig. 68d). In a slightly flattened sporocyte (Fig. 68g-i) the four cones of microtubules emanating from the plastids and bipolar arrays interlinking them are seen. In late meiotic prophase, the bipolar arrays interconnecting the plastids are well developed and chromosomes are distinct (Fig. 68j-1). As the QMS becomes transformed into the spindle, there is a distinct change in [gamma]-tubulin distribution (Fig. 68k) as it becomes less concentrated at the poles and moves into the distal portion of the forming spindle.
The QMS is transformed into a functionally bipolar spindle in prometaphase I by convergence of the four poles in pairs toward the division axis (Fig. 69a-c). In bryopsid mosses, the [gamma]-tubulin moves away from the plastids, which remain fixed at the tetrad poles, and converges on the polar cleavage furrows. In metaphase I (Fig. 69d-f), [gamma]-tubulin is no longer associated with the plastids. It is concentrated at the broad polar regions of the spindle (Fig. 69e). In anaphase I (Fig. 69g-i) the [gamma]-tubulin moves poleward in advance of the chromosomes, then to the proximal surfaces of the reforming nuclei in telophase I (Fig. 69j-1). Microtubules emanating from proximal surfaces of the nuclei and flanking plastids form a phragmoplast array (Fig. 69j).
During a brief inframeiotic interphase (Fig. 70a-c), the QMS is re-assembled from [gamma]-tubulin again associated with the plastids (Fig. 70b). Stages in development of this new microtubule complex (Fig. 70a-f) are virtually indistinguishable from the prophasic QMS (Fig. 68). Interaction of the four cones of microtubules leads to construction of the prophase II spindles. Spindles are organized (Fig. 70g-i) at right angles to each other in the undivided cytoplasm. Metaphase II spindles (Fig. 70j-l) terminate at plastids and [gamma]-tubulin is concentrated at the poles (Fig. 70k). [gamma]-Tubulin organizes phragmoplast microtubules between sister nuclei in telophase II (Fig. 71a-c). [gamma]-Tubulin associated with broad poles of the expanding phragmoplasts (Fig. 71d-f) nucleates secondary phragmoplasts and the tetrad of spores is cleaved according to the polarity established in early meiotic prophase, [gamma]-Tubulin is found in the second division phragmoplasts and is conspicuously absent from the OB in the first division site (Fig. 71e) supporting contention than minimal microtubules are organized in this site during simultaneous cytokinesis. After cleavage of the spore tetrad, [gamma]-tubulin migrates from proximal nuclear surfaces to plastids. In young spores of the tetrad (Fig. 71j-l) a distinct [gamma]-tubulin rich MTOC (Fig. 71k) is associated with the development of a radial system of microtubules (Fig. 71j). TEM shows a remarkably distinct non-membranous body of electron dense particles from which microtubules radiate (Fig. 72). Additional studies of this rare form of corpuscular plant MTOC may provide insight into the functional organization of MTOCs in plant cells.
The Hornworts (Anthocerophytina) are a small (about 500 species in 12 genera) but evolutionarily important group as the most current phylogenies find them to be the closest extant relatives of the tracheophytes (Carafa et al., 2005; Renzaglia et al., 2009; Villarreal et al., 2010). They are unique in that the partially dependent sporophyte has an intercalary meristem at the base of the sporangium (Fig. 73). Archesporial cells are continually produced in this basal meristem and mature spores are released from the slit-like apertures in the apical region of the sporangium. This, at least in some taxa, results in a prolonged period of sporogenesis with successive stages in longitudinal files of cells progressing from mitosis to mature spores. Hornworts have been used as a model system (Brown & Lemmon, 1990, 1993, 1997) for describing the development and function of the QMS in monoplastidic sporogenesis where it can be seen that the nucleus delays division while the cytoplasm, as evidenced by plastid and microtubule behavior, prepares for two divisions and anticipates the eventual cytokinesis into a tetrad. The coordination of plastid and nuclear division involves a bipolar system of microtubules (BMA), a basic component of the cytoskeletal mechanism for establishing domains within a common cytoplasm in preparation for eventual division. For this reason we review here a description of the unusual preparation for division polarity in mitosis of hornworts.
In preparation for mitosis, the single plastid elongates, migrates to a position parallel to the future spindle axis and constricts in the middle. A conspicuous microtubule system assembles at the isthmus and extends to the plastid tips in the polar regions of the cell (Fig. 74a). This system is termed the axial microtubule system (AMS) because of its orientation parallel to the long axis of the plastid and the future spindle. The AMS appears to function both in establishment of division polarity and development of the spindle. The AMS is initiated in advance of the PPB, which subsequently girdles the cell at precisely the midregion of the plastid and predicts the future plane of cytokinesis (Fig. 74a). Division polarity is thus marked sequentially by plastid alignment, assembly of the AMS, and finally by the PPB. Concentration of the [gamma]-tubulin at the plastid tips (Fig. 74b) results in generation of microtubules with plus ends at the equatorial region of the forming spindle (Fig. 74c). The AMS appears to contribute directly to development of the mitotic spindle, which is clearly anchored at the plastid tips (i.e. plastid polarity) and the prophase spindle is decidedly asymmetric, encasing the nucleus from the plastid side (Fig. 74c) before becoming symmetric and forming the two opposing half spindles of prometaphase (Fig. 74d). For additional details of mitosis in hornworts, see earlier discussion of bryophyte mitosis.
In preparation for meiosis, the single plastid elongates, an AMS develops parallel to it and the plastid divides as in mitotic prophase (Brown & Lemmon, 1988b, 1993). Then the two plastids elongate, an AMS forms parallel to each, and the plastids rotate so that they are perpendicular to each other with their tips in tetrahedral arrangement (Brown & Lemmon, 1993). Microtubules emanating from the plastids form additional BMAs interconnecting the plastid tips. The plastids divide and the four resulting plastids serve as MTOCs, from which cones of microtubules forma QMS surrounding the nucleus (Fig. 75a-d). The nucleus becomes quadrilobed (Fig. 75c) and the cytoplasm lobes around the plastid poles with substantial deposition of sporocyte wall material at the boundaries of the opposing microtubule arrays. This results in a quadrilobed sporocyte in which the division planes for quadripartitoning into the spore tetrad are clearly established before the nucleus undergoes division (Fig. 75d).
Sequential stages of sporocyte development are seen in longitudinal files of cells produced by the basal meristem. The final division of the archesporial cells gives rise to daughter cells that depart on different developmental pathways. The basal daughter cell is the pseudoelater mother cell (elaterocyte) and the upper cell is the spore mother cell (sporocyte). Thus, the sporocytes forma developmental series with the oldest stages at the top of the capsule (Figs. 73 and 76a). The sporocytes develop pliable walls, expand rapidly and are released from the archesporial cell walls into the mucilaginous matrix of the sporangium. The early sporocytes have a nucleus surrounded by a sinuous ribbon-like plastid (Fig. 76b). In the early sporocyte no polarity can be recognized in the organization of microtubules (Fig. 77a). The plastid divides and two daughter plastids migrate to opposite sides of the nucleus where they enlarge considerably and accumulate starch (Fig. 76c). In the two-plastid sporocyte, microtubules radiate from along the entire length of the plastids; their plus ends interacting to forma basket around the nucleus (Fig. 77b, c). The plastids develop a central isthmus and rotate so that their tips are tetrahedrally positioned and divide to place the four plastids at the tetrad poles (Figs. 75 and 76d). Microtubules radiating from the plastid surfaces form the QMS, which interconnects the four plastids and surrounds the nucleus that becomes distorted into four points opposite the plastids (Fig. 77d).
The QMS closely envelops the nucleus (Figs. 77d and 78a) and morphs directly into the metaphase I spindle (Fig. 78b). The plastids remain at the tetrad poles (poles of meiosis II) and the relatively small rectangular spindle is suspended from the inner surfaces of pairs of plastids (Fig. 78b). The telophase I nuclei are located between pairs of plastids at the polar cleavage furrows. A phragmoplast develops between them (Fig. 78c) and spreads to the equatorial cleavage furrows (Fig. 78d). However, as is typical of sporogenesis in bryophytes, the phragmoplast does not function in the deposition of a dyad wall and second division spindles develop simultaneously between plastids in the undivided cytoplasm. Second meiosis (not shown) results in each spore domain receiving both a nucleus and plastid. Simultaneous cytokinesis involving a system of phragmoplasts that develops among all four nuclei results in cleavage of the spore tetrad along predetermined planes. The placement and development of the phragmoplasts in simultaneous cytokinesis has been followed in greater detail in other bryophytes.
Acknowledgments We thank the many bryologists who helped with collecting materials and providing identifications. Dr. T. Horio (Univ. Kansas) kindly provided the G9 antibody.
Published online: 24 January 2013
Becker, B. & B. Marin. 2009. Streptophyte algae and the origin of embryophytes. Annals of Botany 103: 999-1004.
Bednara, J. & B. Rodkiewicz. 1984. Distribution of plastids and mitochondria during sporogenesis in Equisetum hyemale, pp 17-20. In: M. T. M. Willemse & J. L. van Went (eds). Sexual reproduction in seed plants, ferns and mosses. Pudoc, Wageningen.
--, L. Gielwanowska & B. Rodkiewicz. 1986. Regular arrangements of mitochondria and plastids during sporogenesis in Equisetum. Protoplasma 130:145-152.
Binarova, E, V. Cenklova, J. Prochazkova, A. Doskocilova, J. Volc, M. Vrlik & L. Bogre. 2006. [gamma]-Tubulin is essential for acentrosomal microtubule nucleation and coordination of late mitotic events in Arabidopsis. Plant Cell 18:1199-1212.
Blackmore, S. & S. H. Barnes. 1987. Embryophyte spore walls: origin, development, and homologies. Cladistics 3: 185-195.
-- & R. B. Knox. 1990. Microspores Evolution and ontogeny. Academic, London.
--, A. H. Wortley, J. J. Skvarla & J. R. Rowley. 2007. Pollen wall development in flowering plants. New Phytologist 174: 483-498.
Brown, R. C. & B. E. Lemmon. 1982a. Ultrastructure of meiosis in the moss Rhynchostegium serrulatum I. Prophasic microtubules and spindle dynamics. Protoplasma 110:23-33.
-- & --. 1982b. Ultrastructure of sporogenesis in the moss. Amblystegium riparium I. meiosis and cytokinesis. American Journal of Botany 69:1096-1107.
-- & --. 1983. Microtubule organization and morphogenesis in young spores of the moss Tetraphis pellucida Hedw. Protoplasma 116:115-124.
-- & --. 1986. Spore wall development in the liverwort, Haplomitrium kookeri. Canadian Journal of Botany 64:1174-1182.
-- & --. 1987a. Division polarity, development and configuration of microtubule arrays in bryophyte meiosis I: meiotic prophase to metaphase I. Protoplasma 137:84-99.
-- & --. 1987b. Division polarity, development and configuration of microtubule arrays in bryophyte meiosis II: anaphase I to the tetrad. Protoplasma 138: 1-10.
-- & --. 1988a. Sporogenesis in bryophytes. Advances in Bryology 3: 159-223.
-- & --. 1988b. Preprophasic microtubule systems and development of the mitotic spindle in hornworts (Bryophyta). Protoplasma 143:11-21.
-- & --. 1988c. Cytokinesis occurs at boundaries of domains delimited by nuclear-based microtubules in sporocytes of Conocephalum conicum (Bryophyta). Cell Motility and the Cytoskeleton 11: 139-146.
-- & --. 1990. Polar organizers mark division axis prior to preprophase band formation in mitosis of the hepatic Reboulia hemisphaerica (Bryophyta). Protoplasma 156:74-81.
-- & --. 1991. The cytokinetic apparatus in meiosis: Control of division plane in the absence of a preprophase band of microtubules, pp 259-272. In: C. W. Lloyd (ed). The cytoskeletal basis of plant growth and form. Academic, New York.
-- & --. 1992. Cytoplasmic domain: a model for spatial control of cytokinesis in reproductive cells of plants. Electron Microscopy Society of America Bulletin 22: 48-53.
-- & --. 1993. Diversity of cell division in simple land plants holds clues to evolution of the mitotic and cytokinetic apparatus in higher plants. Memoirs of the Torrey Botanical Club 25: 45-62.
-- & --. 1995. Methods in plant immunolight microscopy. Methods in Cell Biology 49: 85-107.
-- & --. 1997. The quadripolar microtubule system in lower land plants. Journal of Plant Research 110:93-106.
-- & --. 2001. The cytoskeleton and the spatial control of cytokinesis in the plant life cycle. Protoplasma 215: 35-49.
-- & --. 2004. [gamma]-Tubulin, microtubule arrays, and quadripolarity during sporogenesis in the hepatic Aneura pinguis (L.) Dumort. (Metzgeriales). Journal of Plant Research 117: 371-376.
-- & --. 2006. Polar organizers and girdling bands of microtubules are associated with [gamma]-tubulin and act in establishment of meiotic quadripolarity in the hepatic Aneura pinguis (Bryophyta). Protoplasma 227: 77-85.
-- & --. 2007. The pleiomorphic plant MTOC: an evolutionary perspective. Journal of Integrative Plant Biology 49:1142-1153.
-- & --. 2008. [gamma]-Tubulin and microtubule organization during meiosis in the liverwort Ricciocarpus natans (Ricciaceae). American Journal of Botany 95:664-671.
-- & --. 2009. Pre-meiotic bands and novel meiotic spindle ontogeny in quadrilobed sporocytes of leafy liverworts (Jungermannidae, Bryophyta). Protoplasma 237: 41-49. doi: 10.1007/s00709-009-0073-4.
-- & --. 2011a. Spores before sporophytes: hypothesizing the origin of sporogenesis at the algal-plant transition. New Phytologist 190:875-881. doi:10.1111/j.1469-8137.2011.03709.x.
-- & --. 2011b. Dividing without centrioles: innovative MTOCs organize mitotic spindles in bryophytes, the earliest extant lineages of land plants. AoB Plants 2011. doi:10.1093/aobpla/plr028. & Z. Carothers. 1982a. Spore wall development in Sphagnum lescurii. Canadian Journal of Botany 60: 2394-2409.
-- & --. 1982b. Spore wall ultrastrncture of Sphagnum lescurii Sull. Review of Palaeobotany and Palynology 38: 99-107.
-- & K. S. Renzaglia. 1986. Sporocytic control of spore wall pattern in liverworts. American Journal of Botany 73:593-596.
-- & T. Horio. 2004. [gamma]-Tubulin localization changes from discrete polar organizers to anastral spindles and phragmoplasts in mitosis of Marchantia polymorpha L. Protoplasma 224:187-193.
-- & M. Shimamura. 2007. Transformations of the pleiomorphic plant MTOC during sporogenesis in the hepatic Marchantia polymorpha L. Journal of Integrative Plant Biology 49: 1244-1252.
-- & --. 2010. Diversity in meiotic spindle origin and determination of cytokinetic planes in sporogenesis of complex thalloid liverworts (Marchantiopsida). Journal of Plant Research 123:589-605.
Browning, A. J. & B. E. S. Gunning. 1979a. Structure and function of transfer cells in the sporophyte haustorium of Funaria hygrometrica Hedw. II. Kinetics of labelled sugars and localization of absorbed products by freeze-substitution and autoradiography. Journal of Experimental Botany 30:1247-1264.
-- & --. 1979b. Structure and function of transfer cells in the sporophyte haustorium of Funaria hygrometrica Hedw. II. Translocation of assimilate into the attached sporophyte and along the seta of attached and excised sporophytes. Journal of Experimental Botany 30:1265-1273.
Busby, C. H. & B. E. S. Gunning. 1988a. Establishment of plastid-based quadripolarity in spore mother cells of the moss Funaria hygrometrica. Journal of Cell Science 91: 117-126.
-- & --. 1988b. Development of the quadripolar meiotic cytoskeleton in spore mother cells of the moss Funaria hygrometrica. Journal of Cell Science 91: 127-137.
-- & --. 1989. Development of the quadripolar meiotic apparatus in Funaria spore mother cells: analysis by means of anti-microtubule drug treatments. Journal of Cell Science 93: 267-277.
Carafa, A., J. G. Duckett & R. Ligrone. 2003. The placenta in Monoclea fosteri Hook. and Treubia lacunosa (Col.) Prosk.: insights into placental evolution in liverworts. Annals of Botany 92:299-307.
--, --, J. P. Knox & R. Ligrone. 2005. Distribution of cell-wall xylans in bryophytes and tracheophytes: new insights into basal interrelationships of land plants. New Phytologist 168: 231-240.
Crandall-Stotler, B. 1980. Morphogenetic designs and a theory of bryophyte origins and divergence. BioScience 30: 580-585.
-- & R. E. Stotler. 2000. Morphology and classification of the Marchantiophyta. pp 21-70. In: A. J. Shaw & B. Goffinet (eds). Bryophyte biology. Cambridge University Press, Cambridge.
--, & -- D. G. Long. 2009a. Morphology and classification of the Marchantiophyta. pp 1-54. In: B. Goffinet & A. J. Shaw (eds). Bryophyte biology, ed. 2nd. Cambridge University Press, Cambridge.
--, -- & --. 2009b. Phylogeny and classification of the Marchantiophyta. Edinburgh Journal of Botany 66:155-198.
Crum, H. A. & L. E. Anderson. 1981. Mosses of eastern North America. 2 vols. Columbia University Press, New York.
Davis, B. M. 1899. The spore-mother-cell of Anthoceros. Botanical Gazette 27: 89-109.
Doonan, J. H. & J. G. Duckett. 1988. The bryophyte cytoskeleton. Advances in Bryology 3: 1-31.
--, D. J. Cove, F. M. IC Cork & C. W. Lloyd. 1987. Pre-prophase band of microtubules absent from tip-growing moss filaments, arises in leafy shoots during transition to mtercalary growth. Cell Motility and the Cytoskeleton 7: 138-153.
Duckett, J. G., S. Pressel, K. M. Y. P'Ng & K. S. Renzaglia. 2009. Exploding a myth: the capsule dehiscence mechanism and the function of pseudostomata in Sphagnum. New Phytologist 183: 1053-1063.
Edwards, D., J. G. Duckett & J. B. Richardson. 1995. Hepatic characters in the earliest land plants. Nature 374: 635-636.
Farmer, J. B. 1895. On spore-formation and nuclear division in the Hepaticae. Annals of Botany 9:469-523.
-- 1904. On the interpretation of the quadripolar spindle in the Hepaticae. Botanical Gazette 37:63-65.
Fart, C. H. 1916. Cytokinesis of the pollen-mother-cells of certain dicotyledons. Memoirs of the New York Botanical Garden 6: 253-316.
Finet, C., R. E. Timme, C.F. Delwiche & F. Marletaz. 2010. Multigene phylogeny of the green lineage reveals the origin and diversification of land plants. Current Biology 20: 2217-2222.
Ernest, L. L., E. C. Davis, D. G. Long, B. Cradall-Stotler, A. Clark & M. L. Hollingsworth. 2006. Unraveling the evolutionary history of the liverworts (Marchantiophyta): multiple taxa, genomes and analyses. Bryologist 109:303-334.
Frey, W. & M. Stech. 2005. A morpho-molecular classification of the liverworts (Hepaticophytina, Bryophyta). Nova Hedwigia 81:55-78. Furness, C. A. & P.J. Rudall. 1999. Microsporogenesis in monocotyledons. Annals of Botany 84: 475-499.
--& F B. Sampson. 2002. Evolution of microsporogenesis in angiosperms. International Journal of Plant Science 163:235-260.
Garbary, D. J. & K. S. Renzaglia. 1998. Bryophyte phylogeny and the evolution of land plants: evidence from development and ultrastructure, pp 45-63. In: J. W. Bates, N. W. Ashton, & J. G. Duckett (eds). Bryology for the twenty-first century. Maney and the British Bryological Society, Leeds.
Goffinet, B. & A. J. Shaw. 2009. Bryophyte biology, ed. 2nd. Cambridge, New York.
Graham, L. E. 1993. Origin of land plants. John Wiley, New York.
-- 1996. Green algae to land plants: an evolutionary transition. Journal of Plant Research 109: 241-251.
--, M. E. Cook & J. S. Busse. 2000. The origin of plants: body plan changes contributing to a major evolutionary radiation. Proceedings of the National Academy of Sciences USA 97: 4535-4540.
Gray, J. 1993. Major Paleozoic land plant evolutionary bio-events. Palaeogeograpby, Palaeoclimatology, Palaeoecology 104: 153-169.
Gunning, B. E. S. 1982. The cytokinetic apparams: its development and spatial regulation, pp 229-292. In: C. W. Lloyd (ed). The cytoskeleton in plant growth and development. Academic, London.
Hattori, S. & M. Mizutani. 1959. What is Takakia lepidozioides? Journal of the Hattori Botanical Laboratory 20:285-303.
Hedderson, T. A., R. L. Chapman & W. L. Rootes. 1996. Phylogenetic relationships of bryophytes inferred from nuclear-encoded rRNA gene sequences. Plant Systematics and Evolution 200:213-224.
--, R. Chapman & C. J. Cox. 1998. Bryophytes and the origins and diversification of land plants: new evidence from molecules, pp 65-77. In: J. W. Bates, N. W. Ashton, & J. G. Duckett (eds). Bryology for the twenty-first century. Maney and the British Bryological Society, Leeds.
Heslop-Harrison, J. 1966. Cytoplasmic continuities during spore formation in flowering plants. Endeavour 25:65-72.
-- 1972. Pattern in plant cell walls: morphogenesis in miniature. Proceedings of the Royal Institution of Great Britain 45:335-351.
Horner, H. T., N. R. Lersten & C. C. Bowen. 1966. Spore development in the liverwort Riccardia pinguis. American Journal of Botany 53: 1048-1064.
Juel, H. O. 1897. Die kerntheilungen in den pollenmutterzellen von Hemerocallis fulva und die bei denselben auftretenden unregelmassigkeiten. Jahrbucher fur wissenschaftliche Botanik 30: 205-226.
Kelley, C. B. & W. T. Doyle. 1975. Differentiation of intracapsular cells in the sporophyte of Sphaerocarpos donnellii. American Journal of Botany 62:547-559.
Kenrick, E & C. Crane. 1997. The origin and early evolution of plants on land. Nature 389: 33-39.
Ligrone, R. & R. Gambardella. 1988. The sporophyte-gametophyte junction in bryopbytes. Advances in Bryology 3:225-274.
Liu, B., J. Marc, It. C. Joshi & B. A. Palevitz. 1993. A [gamma]-tubulin-related protein associated with the microtubule arrays of higher plants in a cell cycle-dependent manner. Journal of Cell Science 104: 1217-1228.
Mazia, D. 1961. Mitosis and the physiology of cell division, pp 77-412. In: J. Brachet & A. Mirsky (eds). The cell, vol. 3. Academic, New York.
Miller, N. G. (ed.). 1993. Biology of Sphagnum. Advances in Bryology 5: 1-338.
Mineyuki, Y. 1999. The preprophase band of microtubules: its function as a cytokinetic apparatus in higher plants. International Review of Cytology 187: 1-49.
-- 2007. Plant microtubule studies: past and present. Journal of Plant Research 120:45-51.
Mishler, B. D. & S. P. Churchill. 1984. A cladistic approach to the phylogeny of the "bryophytes". Brittonia 36: 406-424.
Moore, A. C. 1905. Sporogenesis in Pallavacinia. Botanical Gazette 40:81-96.
Neidhart, H. V. 1978. Ultrastructural aspects of sporogenesis in Riella affinis Howe and Underwood (Hepaticae). Journal of Bryology 10:145-154.
Otegui, M. & L. A. Staehelin. 2000. Cytokinesis in flowering plants: more than one way to divide a cell. Current Opinions in Plant Biology 3: 493-502.
Ovenchkina, Y. & B. R. Oakley. 2001. [gamma]-Tubulin in plant cells. Methods in Cell Biology 67:195-212.
Pate, J. S. & B. E. S. Gunning. 1972. Transfer cells. Annual Review of Plant Physiology 23:173-196.
Pickett-Heaps, J. D. 1969. The evolution of the mitotic apparatus: an attempt at comparative ultrastructural cytology in dividing plant cells. Cytobios 1: 257-280.
-- 1974. Cell division in Stichococcus. British Phycological Journal 9: 63-73.
Proskauer, J. 1954. On Sphaerocarpos stipitatus and the genus Sphaerocarpos. Journal of the Linnaean Society of London, Botany 55:143-157.
Qiu, Y.-L., Y. Cho, J. C. Cox & J. D. Palmer. 1998. The gain of three mitochondrial introns identifies liverworts as the earliest land plants. Nature 394: 671-674.
Renzaglia, K. S., R. C. Brown, B. E. Lemmon, J. G. Duckett & R. Ligrone. 1994. Occurrence and phylogenetic significance of monoplastidic meiosis in liverworts. Canadian Journal of Botany 72: 65-72.
--, K. D. McFarland & D. K. Smith. 1997. Anatomy and ultrastructure of the sporophyte of Takakia ceratophylla (Bryophyta). American Journal of Botany 84: 1337-1350.
--, R. J. Duff, D. L. Nickrent & D. J. Garbary. 2000. Vegetative and reproductive innovations of early land plants: implications for a unified phylogeny. Philosophical Transactions of the Royal Society of London B Biological Sciences: 355: 769-793.
--, S. Schuette, R. J. Duff, R. Ligrone, A. J. Shaw, B. D. Mishler & J. G. Duckett. 2007. Bryophyte phylogeny: Advancing the molecular and morphological frontiers Bryologist 110: 179-213.
--, J. C. Villarreal & R. J. Duff. 2009. New insights into morphology, anatomy, and systematics of hornworts, pp 139-171. In. B. Goffinet & A. J. Shaw (eds). Bryophyte biology, ed. 2nd. Cambridge University Press, New York.
Rodkiewicz, B., J. Bednara, A. Mostowska, E. Duda & H. Stobiecka. 1986. The change in disposition of plastids and mitochondria during microsporogenesis and sporogenesis in some higher plants. Acta Botanica Neerlandica 35:209-215.
Rudall, P. J., M. V. Remizowa, A. S. Beer, E. Bradshaw, D. W. Stevenson, T. D. Macfarlane, R. E. Tuckett, S. R. Yadav & D. D. D. Sokoloff. 2008. Comparative ovule and megagametophyte development in Hydatellaceae and water lilies reveal a mosaic of features among the earliest angiosperms. Annals of Botany 101:941-956.
Schepf, E. 1984. Pre- and postmitotic reorientation of microtubule arrays in young Sphagnum leaflets: transitional stages and initiation sites. Protoplasma 120:100-112.
Schmit, A.-C. 2002. Acentrosomal microtubule nucleation in higher plants. International Review of Cytology 220:257-289.
Schuster, R. M. 1984. Evolution, phylogeny and classification of the Hepaticae. pp 892-1070. In: R. M.
Schuster (ed). New manual of bryology, Vol. 2. Hattori Botanical Laboratory, Nichinan.
-- 1992. The Hepaticae and Anthocerotae of North America east of the hundreth meridian, Vol. VI. Field Museum of Natural History, Chicago.
Shaw, J. & K. S. Renzaglia. 2004. Phylogeny and diversification of bryophytes. American Journal of Botany 91: 1557-1581.
Shimamura, M. & H. Deguchi. 2008. Sporophyte anatomy of Oedipodium griffithianum (Oedipodiaceae). pp 319-325. In: I. H. Mohamed, B. B. Baki, A. Nasrulhaq-Boyce, & P. K. Y. Lee (eds). Bryology in the new millennium. University of Malaysia Press, Kuala Lumpur.
--, & Y. Mineyuki. 1998. Meiotic cytokinetic apparatus in the formation of the linear spore tetrads of Conocephalum japonicum (Bryophyta). Planta 206:604-610.
--, Y. Mineyuki & H. Deguchi. 2000. Monoplastidic meiosis in Dumortiera hirsuta (Bryophyta; Marchantiales). Journal of the Hattori Botanical Laboratory 88:267-270.
--, A. Kitamura, Y. Mineyuki & H. Deguchi. 2001. Occurrence of monoplastidic sporocytes and quadripolar microtubule systems in Marchantiales (Bryophyta; Marchantiidae). Hikobia 13:551-562.
--, It. Deguchi & Y. Mineyuki. 2003. A review of the occurrence of monoplastidic meiosis in liverworts. Journal of the Hattori Botanical Laboratory 94: 179-186.
--, R. C. Brown, B. E. Lemmon, T. Akashi, K. Mizuno, N. Nishihara, K.-I. Tomizawa, K. Yoshimoto, H. Deguchi, H. Hosoya, T. Horio & Y. Mineyuki. 2004. [gamma]-Tubulin in basal land plants: Characterization, localization and implication in the evolution of acentriolar microtubule organizing centers. Plant Cell 16:45-59.
--, T. Furuki & H. Deguchi. 2005. Sporophyte anatomy of Cavicularia densa (Blasiaceae). Bryologist 108: 420-426.
--, M. Itouga & H. Tsubota. 2011. Evolution of apolar sporocytes in marchantialean liverworts: implications from molecular phylogeny. Journal of Plant Research. doi: 10.1007/s10265-0425-y. Smith, D. K. 1990. Sporophyte of Takakia discovered. Bryological Times 57(58): 1-4.
Taylor, W. A. & P. K. Strother. 2008. Ultrastructure of some Cambrian palynomorphs from the Bright Angel Shale, Arizona, USA. Review of Palaeobotany and Palynology 151:41-50.
Tobe, H., Y. Kimoto & N. Prakash. 2007. Development and structure of the female gametophyte in Austrobaileya scandens (Austrobaileyaceae). Journal of Plant Research. 120:431-436.
Turner, R. G. 1993. Peat and people: a review. Advances in Bryology 5:315-328.
Villarreal, J. C., D. C. Cargill, A. Hagborg, L. Soderstrom & K. S. Renzaglia. 2010. A synthesis of hornwort diversity: Patterns, causes and future work. Phytotaxa 9: 150-166.
Von Konrat, M., J. Shaw & K. S. Renzaglia. 2010a. A special issue of Phytotaxa dedicated to Bryophytes: The closest living relatives of early land plants. Phytotaxa 9:5-10.
--, L. Soderstrom, M. A. M. Renner, A. Hagborg, L. Briscoe & J. J. Engel. 2010b. Early land plants today (ELPT): How many liverwort species are there? Phytotaxa 9: 22-40.
Wallace, S., A. Fleming, C. H. Wellman & D. J. Beerling. 2011. Evolutionary development of the plant spore and pollen wall. AoB Plants. doi:10.1093/aobpla/plr027.
Wasteneys, G. O. 2002. Microtubule organization in the green kingdom: chaos or self-order? Journal of Cell Science 115:1345-1354.
Wiese, C. & Y. Zheng. 2006. Microtubule nucleation: [gamma]-tubulin and beyond. Journal of Cell Science 119: 4143-4153.
Williams, J. H. & W. E. Friedman. 2004. The four-celled female gametophyte of Illicium (Illiciaceae; Austrobaileyales): implications for understanding the origin and early evolution of monocots, eumagnoliids, and eudicots. American Journal of Botany 91 : 332-351.
Roy C. Brown (1,2), Betty E. Lemmon (1)
(1) Department of Biology, University of Louisiana-Lafayette, Lafayette, LA 70504, USA
(2) Author for Correspondence; e-mail: firstname.lastname@example.org
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|Title Annotation:||p. 248-280|
|Author:||Brown, Roy C.; Lemmon, Betty E.|
|Publication:||The Botanical Review|
|Date:||Jun 1, 2013|
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