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Anther Plastids in Angiosperms.

I. Abstract

In the anther of angiosperms, all types of plastids are found in the course of pollen development. They are located in the different cell layers of the microsporangium and have various functions that contribute to the formation of the functional male gametophyte. This includes photosynthesis, stomata opening, sugar storage and/or mobilization, lipid synthesis and secretion for pollenkitt formation, as well as serving as a physiological buffer under stress conditions. They are also involved in plastid inheritance, but to different extents, according to the species.

The plastid is a semi-autonomous organelle. Plastid division in the anther is synchronous with cell division, except in the vegetative cell during pollen maturation. Furthermore, recent data seem to show that plastids are affected by programmed cell death and DNA degradation, which occur in the whole anther throughout pollen development. However, the timing of plastid disappearance fluctuates in the different cell layers and also depending on species.

In vitro, following androgenesis, plastids that originate in the microspore are responsible for the occurrence of albino plantlets in Poaceae. This trait reflects the relative independence of the plastid genome when compared with that of the nucleus. In this family, microspore plastids may become so involved in programmed cell death that they are unable to follow the alternative sporopohytic program.

The different pathways of plastid differentiation in neighboring anther cell layers require an accurate regulation of cell development that remains widely unknown in the anther.

Resume

Dans l'anthere des angiospermes, on trouve tous les types de plastes au cours du developpement pollinique. Ils sont localises dans les differents couches cellulaires du microsporange et possedent des fonctions variees contribuant la formation du gametophyte male fonctionnel. Cela recouvre la photosynthese, l'ouverture des stomates, le stockage et/ou la mobilisation de glucides, la synthese et la secretion de lipides pour la formation du pollenkitt, ainsi qu'un role de tampon physiologique en conditions stressantes. Ils sont egalement impliques dans l'heredite plastidiale a un degre divers en fonction des especes.

Le plaste est un organite semi-autonome. La division plastidiale au sein de l'anthere est synchrone avec la division cellulaire, excepte dans la cellule vegetative pendant la maturation du pollen. Par ailleurs, des donnees recentes semblent montrer que les plastes sont affectes par la mort cellullaire programmee et la degradation d'ADN qui se produisent dans la totalite de l'anthere tout au long du developpement pollinique. Toutefois, la chronologie de disparition des plastes fluctue dans les differentes couches cellulaires et aussi en fonction des especes.

In vitro, durant le processus d'androgenese, les plastes originaires de la microspore sont responsables de l'apparition de plantules albinos chez les Poaceae. Ce caractere reflete la relative independance du genome plastidial par rapport au genome nucleaire. Dans cette famille, il est probable que les plastes de la microspore sont trop engages dans le programme de mort cellulaire et demeurent incapables de suivre un nouveau programme sporophytique.

Les differentes voies de differenciation plastidiale observees dans les couches cellulaires voisines de l'anthere requierent une regulation precise du developpement cellulaire qui demeure largement inconnue dans l'anthere.

II. Introduction

Plant-cell differentiation is a process in which many cell compartments are involved. Among these, plastids play a crucial role. Meristematic cells exclusively include proplastids. In differentiated cells, plastids can be involved in anabolism (chloroplasts) or in storing reserves in amyloplasts and elaioplasts (Wheatley, 1977). The final phase of differentiation is often characterized by the presence of chromoplasts in various tissues, including senescent leaves, fruit pulp, and petals (Ljubesic et al., 1991). In the vegetative organs, plastid differentiation is usually unidirectional, whereas in the reproductive cells plastid interconversion is frequent (Pacini et al., 1992a).

In angiosperms, the anther comprises several cell types that are involved in supporting the development and nourishment of the pollen grain (Fig. 1). All of the types of plastids can be found in the anther, according to the cell type and the stage of development (Fig. 2). Before meiosis, all of the anther cells have proplastids, and in some cases they may accumulate starch. This was observed in several species and in different cell types; for example, in the tapetal cells of Zea mays (Panchaksharappa & Rudramuniyappa, 1974). As soon as meiosis starts, plastids in the meiocytes are commonly undifferentiated (Dickinson, 1981), whereas in other cell types, plastids can still produce or store some substances. Previous research on anther plastids focused mainly either on tapetum or microspore pollen grains (Pacini et al., 1992a; Pacini, 1996; Clement et al., 1998) or on plastid inheritance (Hageman & Shroder, 1989; Corriveau et al., 1990; Sodmergen et al., 1992, 1994, 1998; Nagata et al., 1999a, 1999b).

The aim of this article is to give an overview of plastid diversity, differentiation, and roles in the anther, with respect to the completion of pollen development and functionality in the reproductive process of angiosperms. In fact, the plastid function of some cells is to support the formation of other cells--for example, the tapetum and the vegetative cell--whereas others, especially sperm cells, sometimes contain plastids responsible for biparental cytoplasmic inheritance and thus are directly involved in the reproductive process. Furthermore, plastids of the microspore/pollen grains interfere with the success of androgenesis, especially in Poaceae. Indeed, in this family a number of microspore-derived androgenetic haploid plantlets are albinos, and the plastids of the initial microspore are involved in this process (Caredda et al., 2000). Therefore, the data presented here will deal with anther plastids during both in vivo and in vitro development.

III. The Anther Wall

A. THE CONNECTIVE TISSUE

The connective tissue is formed of cells that contain undifferentiated plastids during the whole life of the anther under natural conditions (Fig. 3). Besides, plastids in this area can accumulate starch under stress conditions (Saini, 1997; Saini & Lalonde, 1998). In Triticum aestivum, when the whole plant undergoes water stress, the anther walls, especially the connective tissue, accumulate large amounts of starch, indicating an inhibition of sucrose utilization in the anther (Lalonde et al., 1997). In Lilium, when anthers develop in vitro using large amounts of sucrose in the medium, plastids of the connective tissue fill with starch, whereas pollen sugar metabolism is unaffected. This means that the connective tissue acts as a carbohydrate buffer for developing pollen grains (Clement & Audran, 1995).

B. THE LOCULE SURROUNDING EXTERNAL LAYERS: EPIDERMIS, ENDOTHECIUM, AND MIDDLE LAYERS

In these cell layers, plastids are mainly involved in sugar physiology, either storing carbohydrates in starch grains or performing photosynthesis during early pollen development (Clement & Audran, 1999).

Until meiosis, plastids in the outer anther wall are proplastids. Afterward, one or two waves of amylogenesis/amylolysis may occur during development. In the former case, starch accumulates at the beginning of microspore vacuolation and is mobilized during microsporogenesis (Biddle, 1979; Pacini & Franchi, 1983; Bhadula & Sawhney, 1989), from the first haploid mitosis (Keijzer & Willemse, 1958a), or progressively during pollen development (Hourcade et al., 1986). In the latter case, the first cycle of starch synthesis/degradation takes place during microspore vacuolation, whereas the second one occurs around pollen mitosis (Reznickova, 1978; Reznickova & Willemse, 1980; Bhadula & Sawhney, 1989; Clement et al., 1994). In this case, the first wave of starch degradation can be correlated with parallel anther growth, formation of endothecium thickenings, and tapetum development, whereas the second may be due to the arrest of anther growth and a subsequent surplus of soluble sugars from the filament (Clement et a l., 1996a). This may also occur when the blooming period extends for a long time and the plant receives huge amounts of light, producing available nutrients without any limitation (Bellani et al., l985a, 1985b).

Chloroplasts are the most common type of plastid in vegetative organs. They are not as common in reproductive cells, although they are present in the anther wall (Figs. 2B, 2D), especially in the epidermis, the endothecium, and the middle layers throughout microsporogenesis (Kirichenko et al., 1977; Mlodzianowski & Idzikowska, 1978; Gori, 1982; Keijzer & Willemse, l988a, 1988b; Clement et al., 1997). It has been demonstrated that net positive photosynthesis is transiently performed in these chloroplasts, which thus participate in the carbonaceous nutrition of the anther. In Triticum aestivum, in Secale cereale (Kirichenko et al., 1992, 1993) and in Lilium (Clement et al., 1997), anther-wall chloroplasts contain large amounts of chlorophyll compatible with photosynthesis, and the rate of [CO.sub.2] fixation is higher than in the corresponding leaves considering the chlorophyll content. Later on, from pollen mitosis, chloroplasts further develop into chromoplasts by accumulating plastoglobules in the stroma (F ig. 2A), in parallel with the loss of chlorophyll and photosynthetic membranes (Clement et al., 1997).

Plastids of the locule external surrounding layers then cover two important functions regarding pollen development (Fig. 4): they are the favorite place for starch accumulation in the anther, and they are involved in anther carbohydrate production, performing photosynthesis. These cell layers also participate in the buffer function of the anther wall, together with the connective tissue by storing the surplus of soluble sugars in starch (Clement & Audran, 1995).

Stomata are not present in all anthers. But when they are, they can face the connective tissue, as Keijzer (1987) found, or all around the theca, as in Lilium (Clement et al., 1997). Plastids in the anther stomata are chloro-amyloplasts (Clement et al., 1994), acting in the opening of guard cells during the photosynthetic process or acting during anther dehydration before they open (Keijzer, 1987).

Data are still lacking concerning plastids in the outer anther wall, although they may play important functions, especially by synthesizing carbohydrates for internal tapetum and pollen grains, as well as controlling the amount of sugars reaching the developing pollen grains.

IV. The Tapetum

Elaioplast is the final stage of tapetal plastid development (Fig. 2E), but this is reached in different ways according to the species with respect to the onset of the process (Fig. 5). In most cases, tapetal proplastids undergo division during the early stages of microsporogenesis and next develop into elaioplasts (Dickinson, 1973; Pacini & Juniper, 1979b; Keijzer & Willemse, 1988a; Murgia et al., 1991; Pacini et al., 1992a; Weber, 1992, 1996; Giampolini et al., 1993; Hesse & Hess, 1993; Hess & Hesse, 1994; Clement et al., 1998). Then plastids have three possible fates (Pacini, 1997), mainly according to the mode of pollination.

First, in strictly anemophilous species, such as Lolium perenne, plastids first develop into chromoplast, later in elaioplast, and then are resorbed with the other tapetal cell components (Pacini et al., 1992a). In Lycopersicon peruvianum, another anemophilous species, the pathway is the same but without the chromoplast stage.

Second, in entomophilous species, elaioplasts produce large amounts of lipids in their stroma, which are released into the cytoplasm through the plastid envelopes prior to tapetal plasma membrane disruption (Heslop-Harrison, 1968; Dickinson & Lewis, 1973; Pacini & Juniper, 1979b; Reznickova & Willemse, 1980, 1981; Reznickova & Dickinson, 1982; Keijzer & Willemse, 1988b; Murgia et al., 1991; Pacini et al., 1992a; Weber, 1992, 1996; Hesse & Hess, 1993; Hess & Hesse, 1994). Moreover, during microsporogenesis elaioplasts cooperate with the endoplasmic reticulum to produce the lipids of the spherosomes (Ciampolini et al., 1993; Hesse, 1993; Hess & Hesse, 1994; Clement et al., 1998) by in situ degeneration (Pacini & Franchi, 1991). Finally, prior to tapetum degeneration, elaioplast lipids mix with spherosomes and constitute the main part of the pollenkitt, which will reach the pollen wall (Fitzgerald et al., 1994).

Third, the Brassicaceae represent a special case, because it is an entomophilous family. Both tapetosomes and elaioplasts are involved in the formation of tapetal lipids that are released in the locule to form tryphine (Dickinson & Lewis, 1973) by extra situm degeneration (Pacini & Franchi, 1991). Tapetasomes include oleosins and triacylglycerols, whereas elaioplasts contain neutral esters, which represent the major lipid component of the pollen coat (Ting et al., 1998). Elaioplasts are released into the loculus simultaneously with the other elements of the cytoplasm when the tapetal plasma membrane breaks down (Dickinson & Lewis, 1973). The tapetum cell components then reach the wall of the developing pollen grains, and tryphines are formed (Pifanelli et al., 1998).

In a few cases, including Pisum sativum (Biddle, 1979),Austrobaileya maculata (Zavada, 1984), Trillium kamtshaticum (Takahashi, 1987), Gasteria verrucosa (Keijzer & Willemse, 1988a, 1988b), Brassica oleracea (Murgia et al., 1991), Arachis hypogea (Xi, 1991) and Tilia platyphyllos (Hesse, 1993), tapetal plastids transiently store starch at the beginning of microsporogenesis. These reserves are mobilized during microspore vacuolation and completely disappear when the microspore has completed the vacuolation process.

Even in the same family-for example, the Solanaceae-the pathway of plastid development in the tapetum is the same until elaioplast differentiation. Later on, it varies according to whether pollination is anemophilous (Lycopersicon peruvianum) or entomophilous (Petunia hybrida) (Pacini, unpubl. data). In fact, in the latter case, pollenkitt is necessary to stick the pollen grains together and also to the pollinator body.

V. The Meiocyte/Microspore/Pollen Grain

A. PLASTID AUTONOMY AT MEIOSIS

Meiosis is not only the passage from the sporophyte to the gametophyte phase, where the main compartment of the cell involved is the nucleus. Semi-autonomous organelles, such as plastids and mitochondria, also become involved. In Lilium meiocytes, it was demonstrated that DNA is synthesized in the plastids independently of surrounding meiosis and cytoplasm clearing (Dickinson, 1981; Bird et al., 1983). At the end of meiosis, plastids are then prepared to undergo the whole pollen development (Fig. 6).

B. PLASTIDS: A STORAGE SITE FOR NUTRIENTS AND GROWTH PROMOTERS

Microspore/pollen plastids are mostly known to act as a storage place for carbohydrate reserves, although they are involved in other fundamental processes, such as the promotion of pollen-tube growth.

From meiocytes to ripe pollen grain, only proplastids (Fig. 2A) and amyloplasts (Fig. 2F) can be found in the pollen of angiosperm species. During development, one or two cycles of amylogenesis/amylolysis occur, according to the species. Species with one wave are the dicots Gossypium hirsutum (Wetzel & Jensen, 1992) and Cucurbita pepo (Nepi et al., 1996) and the monocots Ophrys lutea (Feijo & Pais, 1988), Tillandsia pallidoflavescens (Hess, 1991), Lolium perenne (Pacini et al., 1992b), and Lilium hyb. Enchantment (Clement et al., 1994). Species with two waves are the dicots Oenothera (Noher de Halac et al., 1990), Nicotiana tabacum (Olmedilla et al., 1991), Parietaria judaica, Prunus avium, and Lycopersicum peruvianum (Franchi & Pacini, 1988) and the monocots Similax aspera (Pacini & Franchi, 1983) and Pterostylis plumosa (Pandolfi et al., 1993). In most species, the occurrence of amylogenesis in the developing microspore/pollen grain is always preceded by vacuolation (Pacini, 1994). This suggests that starc h synthesis/degradation may be involved in the vacuolation process, probably by regulating the osmotic pressure in the vacuolar sap providing or storing osmoticum as glucose monomers.

With respect to plastids, ripe pollen grains can be divided into two categories: with amyloplasts and with proplastids. If starch is not present, it does not mean that carbohydrate reserves are absent. But these are localized in the cytoplasm of both the vegetative and the generative cell (Franchi et al., 1996). With respect to carbohydrate localization, we can distinguish three categories: only with starch; with starch and cytoplasmic carbohydrates; and only with cytoplasmic carbohydrates (Franchi et al., 1996; Speranza et al., 1997). The more common categories are the second and the third and correspond to those having longer life.

In addition to providing nutrients for energy or monomers for the construction of the pollen-tube wall, plastids of the vegetative cell are involved in pollen-tube growth. Indeed, it has been demonstrated that pollen plastids of Lolium perenne store brassinosteroids during maturation and thus promote pollen-tube growth after pollination (Taylor et al., 1993).

C. SYNCHRONY IN PLASTID DIFFERENTIATION

The anther tapetal cells behave synchronously, whereas mejocytes/microspores/pollen grains lose their synchrony as early as the second meiotic division because cytomictic channels are sealed off (Pacini & Juniper, 1984). Owing to this asynchrony, some microspores start their process of differentiation earlier than do others. Massulatae orchids also behave synchronously in this respect, because the cytomictic channel persists until the early bicellular stage (Heslop-Harrison, 1968). The most noticeable correlated event we have noted is the asynchronous appearance of amyloplast in microspores in the loculus. In contrast, anther dehydration, which interrupts the process of development of the pollen grain, is a synchronous process. That is why, in some species, ripe pollen may or may not have starch (Franchi et al., 1996).

When plastid starts to accumulate starch, one or many granules may be synthesized (Pacini. 1996). This probably has a metabolic meaning, in the sense that the higher the surface of starch grains per plastid the faster the mobilization of starch.

Plastid differentiation is a synchronous process in each cell; that is, all the plastids of a cell differentiate and dedifferentiate simultaneously. There is an exception concerning amyloplast formation during the bicellular stage of some grasses, such as Sorghum bicolor, in which the plastids next to the tapetum accumulate starch before those that are far from the tapetum do (Christensen & Homer, 1974). On the contrary, in another grass, Lolium perenne, all the plastids of the vegetative cell accumulate starch synchronously (Pacini et al., 1992b).

D. POLLEN PLASTIDS AND PROGRAMMED CELL DEATH

Progammed cell death naturally occurs in the anther during in vivo pollen development (Wang et al., 1999a). In the vegetative cell of pollen grain, plastid DNA breaks down from pollen mitosis independently of maternal or biparental inheritance (Sodmergen et al., 1992). Nevertheless, in Lilium longiflorum, a species with maternal inheritance, nucleases are active in the ripe pollen grain, whereas they are not present in Pelargonium zonale, a species with biparental inheritance. In the latter species, the disappearance of plastid DNA coincides with the synthesis of starch in the vegetative cell (Sodmergen et al., 1994), suggesting a correlation between starch metabolism and plastid differentiation in the pollen grain.

In the microspore/pollen grain, the degradation of plastid DNA takes place in the vegetative cell after pollen mitosis in all investigated species (Nagata et al., 1999a, 1999b). However, fluctuations were observed in the timing of plastid DNA degradation during the stages of pollen development, according to the species or to cultivars in the same species. In Hordeum vulgare, the winter cv. Igri and the spring cv. Cork behave in an opposite manner. In the former, 15.3 [+ or -] 2.70% of the microspore plastids contain DNA; in the latter, only 1.7 [+ or -] 0.47% do (Caredda et al., 2000). This has a direct effect on the yield of androgenesis, especially the rate of albino regenerated plantlets.

E. POLLEN AMYLOPLASTS AND ALLERGY

During the last few years, some researches have demonstrated that some allergenic proteins, such as Lol p 5, are localized on starch granules of ripe grass pollen grains (Knox & Suphioglu, 1996). During summer thunderstorms, these pollen grains burst because of water and release large amounts of 3-10-micrometer starch granules, which induce asthmatic attacks when they are breathed in by sensitive persons. Interestingly, the appearance of asthma symptoms in patients is correlated with the appearance of starch in the pollen grains (Linskens et al., 1980), meaning that the proteins responsible for allergic attacks may be involved in pollen starch metabolism.

VI. Plastid Division

Plastid division is a typical feature of cell differentiation (Wheatley, 1977). The plastids of the whole anther cell types divide during the premeiotic stages; later, those of the anther walls differentiate but no longer divide (Pacini et al., 1992a). Those of the tapetum and meiocytes-microspore-vegetative cell, instead, divide again before storing substances, essentially lipids and starch, respectively.

In the tapetum, plastid divisions may occur either before lipid accumulation, as in Cucurbita (Ciampolini et al., 1993), or when elaioplasts are already differentiated, as in Lilium (Clement et al., 1998). Plastid division before reserve deposition is necessary to create many sites in which to store and mobilize a large number of substances in a reduced time. Elaioplasts have two possible fates: to fuse together and with spherosomes to produce the pollenkitt, or to be resorbed, together with all of the other tapetal cell remnants (Pacini et al., 1992a). Even in the latter case, the large number of elaioplasts per tapetal cell ensures rapid mobilization.

In microspore/pollen grains the density of plastids in the cytoplasm varies during pollen development. In Hordeum vulgare, plastids are scarce in the microspore mother cell (2.8 [+ or -] 0.3 plastids per 100 [micro][m.sup.2] cytoplasm), and they intensely multiply later on, during early vacuolation, reaching up to 72.3 [+ or -] 2.4 plastids per 100 [micro][m.sup.2] cytoplasm before the first pollen mitosis (Caredda et al., 2000). In Lilium, this wave of plastid divisions begins earlier, because synthesis of plastid DNA takes place at the premeiotic stage (Dickinson, 1981; Bird et al., 1983).

Plastid division is usually closely correlated with cell division (Pyke, 1997). However, in some species amyloplasts are still able to divide during late pollen maturation, even though the pollen mitosis (es) is (are) achieved, This occurs when vegetative cell plastids of Lolium perenne or Lilium pollen store a starch grain at one pole and this is successively pinched off (Pacini et al., 1992a; Clement et al., 1996a). The process continues until anthesis, which results in the wide increase of amyloplasts in the vegetative cell. This is of importance regarding the fate of starch reserves: the higher the number of amyloplasts and the higher the surface of starch grains, the faster the mobilization that may occur before shedding or during germination. This should be correlated with the fact that in some pollen grains amyloplasts contain only one starch grain, as in Lolium and other grasses (Pacini et al., 1992b), and in others, some hundreds, as in Magnolia soulangeana and other Magnoliaceae (Pacini, unpubl. data).

VII. The Generative Cell and Plastid Inheritance

Plastid inheritance in angiosperms can be paternal, maternal, or biparental, although maternal inheritance seems to be more widespread than biparental (Hageman & Schroder, 1989; Mogensen, 1996).

A. PLASTID TRANSMISSION

In maternal inheritance, male plastids do not participate in the zygote, because they are excluded during pollen development (early) or pollen-tube growth (late). In early plastid exclusion, there is either a priori exclusion of plastids from the generative cell at the time of first haploid mitosis or a posteriori exclusion afterward (Pacini et al., 1992a).A priori exclusion is performed by unequal organelle distribution in the microspore before mitosis. The components mostly invoked are the cytoskeleton, including microfilaments (Schroder, 1985; Hageman & Schroder, 1989) and microtubules (Tanaka, 1991; Mogensen, 1996) gathering plastids apart from the generative cell, as well as the endoplasmic reticulum (ER) network forbidding the plastids to join the cytoplasm of the generative cell (Pacini, 1994). A posteriori exclusion of plastids occurs either during pollen maturation or during pollen-tube growth by degeneration (Pacini et al., 1992a). Late plastid exclusion is performed at the time of fertilization, a s in Triticum and Triticale: it seems that plastids are separated from the sperm nucleus just before penetration (Hageman & Schroder, 1989), but the process remains unclear in most species.

Biparental inheritance is very rare in angiosperms and has been described in only a few species--namely, Medicago sativa (Schumann & Hancock, 1989; Corriveau et al., 1990), Daucus (Hause, 1991), Pelargonium zonale (Sodmergen et al., 1992, 1994, 1998), Rhododendron mucronatum (Nagata et al., 1999a), and Pharbitis nil (Nagata et al., 1999b)--without any report on the mechanism of plastid distribution during pollen development. Plastids are incorporated, remain in the generative cell, and participate in the formation of the zygote (Hageman & Schroder, 1989; Mogensen, 1996; Nagata et al., 1999a). Plumbago zeylanica is a particular case of biparental inheritance. Actually, during the second haploid mitosis, one sperm cell is provided exclusively with mitochondria, whereas the other one contains only plastids. The mitochondria-containing sperm cell fuses with the central cell, and the plastid-containing sperm cell fuses with the egg cell (Mogensen, 1996), causing maternal inheritance for mitochondria and biparenta l inheritance for plastids.

In sperm cells, the health of surviving plastids is often correlated with their DNA content. Indeed, in maternal inheritance with a posteriori exclusion, plastid DNA is degraded after the first pollen mitosis during pollen maturation (Miyamura et al., 1987; Nagata et al., 1 999a). In species with biparental inheritance, however, plastid DNA is intensively synthesized during the end of pollen maturation (Miyamura et al., 1987), and DNA content is multiplied up to twelvefold (Nagata et al., 1999b). Plastid DNA is thus preserved until fertilization (Sodmergen et al., 1992, 1994, 1998) and can be transmitted to the next generation (Mogensen, 1996). Furthermore, plastids and mitochondria are independent in terms of inheritance. In the sperm cells, DNA strongly increases in the organelles that will belong to the zygote, whereas the DNA progressively cdisappears in organelles that are not involved in cytoplasmic inheritance (Nagata et al., 1999a, l999b). The four intermediate cases (mt+/pt+, mt+/pt-; mt-/pt+, mt-/p t-) were observed in angiosperms (Nagata et al., 1999a).

B. PLASTID SEGREGATION AT THE FIRST HAPLOID MITOSIS

Independently of maternal or biparental inheritance, the degree of plastid differentiation may interfere with the entrance of plastids into the generative cell. When the first haploid mitosis occurs, plastids in the microspore can be proplastids or amyloplasts. If they enter the generative cell, they are always in the proplastid state (Pacini et al., l992a). This seems to indicate that they must be undifferentiated to be included in the generative cell. Later on, during pollen maturation, the fate of plastids in the vegetative and generative cells diverges. In fact, those in the generative cell remain proplastids, whereas those in the vegetative cell may differentiate into amyloplasts (see above). Lolium perenne is the only exception to this rule: plastids in the generative cell accumulate starch concurrently with those in the vegetative cell, but they are successively degraded, and inheritance is of the maternal type (Pacini et al., 1992b).

VIII. Genetic and Environmental Influences on Starch Accumulation in Pollen

Plastid differentiation/dedifferentiation in pollen is under genetic and/or environmental control. In Zea mays, the genome seems to influence the presence or absence of starch in pollen grains. Indeed, the segregation of genes can be responsible for the presence of starch in ripe pollen grains, for 50% of the pollen is starchless and 50% is starchy (Hixon & Brimhall, 1968). Also, in Mercurialis annua, a species that blooms year-round, pollen plastids are always devoid of starch (Lisci et al., 1994). However, in other species, such as Parietaria judaica, which blooms from April to October, the percentage of pollen with starch grains progressively decreases from spring to autumn, suggesting the involvement of physiological conditions due to the environment in plastid differentiation (Franchi et al., 1984).

Both the environment and genomes may influence plastid differentiation in pollen grains. In species with cleistogamy, pollen grains from cleistogamous flowers contain starch, whereas those of chasmogamous flowers are devoid of it (Franchi et al., 1996). The appearance of cleistogamous flowers is triggered by adverse environmental conditions, whereas chasmogamous flowers bloom under optimal conditions (Lord, 1981). It should be noted that starchless pollen grains survive longer than do those containing starch (Franchi et al., 1996).

IX. Microspore Plastids and Androgenesis

Androgenesis consists of regenerating in vitro haploid plantlets from unicellular microspores that are initially programmed to develop into a pollen grain. The switch from the gametophytic program to the alternative sporophytic program is initiated using different types of stress, inducing the embryogenic development of microspores (Garrido et al., 1995). Two aspects of androgenesis

may be linked to the microspore plastids.

A. STARCH GRAINS AND MICROSPORE COMPETENCE IN ANDROGENESIS

The uninucleate microspore stage is uniformly recognized as the most suitable stage of pollen development for initiating androgenesis with greatest chances of success. However, it was demonstrated that proplastid-containing microspores usually enter embryogenesis, whereas starch-containing microspores are unable to switch their development toward the sporophytic program (Sangwan & Sangwan-Norreel, 1987; Kott et al., 1988; Zaki & Dickinson, 1990). In the microspore, the degree of plastid differentiation can then be correlated with microspore competence to modify its genetic program. Therefore, starch appears as a marker of irreversible cell differentiation in the microspore. This was confirmed in Zea mays: starch synthesis in microspore plastids coincided with the appearance of new proteins, suggesting a modification in the genome expression, and, concurrently, the microspores lost their androgenetic potential (Mandaron et al., 1990).

B. ALTERATION OF MICROSPORE PLASTID DNA IN POACEAE

In the single Poaceae family, all of the species tested for androgenesis give rise to a certain proportion (from 3 to 100%) of albino microspore-derived plantlets (Caredda & Clement, 1999). This decreases the potential use of androgenesis in these plants of great economic interest (Jahne & Lorz, 1995). Plastids are present in microspore-derived structure during the whole androgenetic process (Sunderland & Huang, 1985). Later on, when androgenetic embryos regenerate plantlets, plastids develop either into chloroplasts (chlorophyllous pathway) or albino plastids (albino pathway). In the latter, it has been shown that plastid DNA is altered. In Hordeum vulgare, transcripts for the plastid genes rbcL, psbD-psbC and the 16S and 23S plastid RNAs are less represented than they are in chlorophyllous plantlets. Moreover, the nuclear genome is also affected. Actually, the nuclear genes rbcS and Cab (encoding for plastiddestined proteins) are underexpressed in albino plantlets. In Oryza sativa (Sun et al., 1979) and in Triticum aestivum (Day & Ellis, 1984, 1985; Ellis & Day, 1986), similar modifications of plastid DNA were observed.

The nature of pretreatment that induces the modification of the microspore program toward the alternative sporophytic program influences the fate of microspore plastids in the pollen embryo. In this respect, the cold pretreatment (4[degrees]C for 28 days) generates irreversible alterations of plastids, increasing the proportion of albino regenerated androgenetic plantlets to a large extent. In contrast, mannitol pretreatment (30 g/l for 3 days) better preserves plastid structure and function, leading to higher rates of chlorophyllous plantlets (Caredda et al., 1999).

DNA defects in both plastids and nuclei of androgenetic plantlets may originate as early as the microspore. Actually, Poaceae belong to a species with cytoplasmic maternal inheritance (Hageman & Schroder, 1989), meaning that semi-autonomous organelles within both the vegetative and the generative cells are programmed to disappear at the end of pollen-tube germination. This suggests a progressive degradation of DNA during pollen development (Mogensen, 1996), which may explain the occurrence of albino microspore-derived plantlets. Recently, Wang et al. (1999a) showed that cell death is programmed in the anther of Hordeum vulgare. DNA is progressively degraded in all anther cell types during in vivo pollen development. During the androgenetic process, the culture conditions stop the programmed cell death and restore DNA defections before entering the embryogenic process (Wang et al., 1999b). In albino microspore-derived plantlets, it is probable that the restoration is not fully achieved, especially in the plas tids. In this species it also seems that plastid DNA degradation may occur in the microspore prior to sampling, particularly in spring cultivars known to generate up to 99% of albino plantlets following androgenesis (Caredda et al., 2000).

X. Conclusions

Anther plastids directly or indirectly cover multiple functions related to the achievement of sexual reproduction (Fig. 7). They are involved in: pollen development, including nutrition, vacuolation by controlling the osmotic pressure, and regulation of carbohydrate supply; pollination, such as pollinator attraction, pollen adherence to pollinator, and pollen protection against UV radiations; pollen survival, by participating in regulating the sugar content of the cytoplasm; and pollen germination, by adjusting the osmotic pressure and providing energy and material for pollen-tube growth.

The different plastid pathways probably arise from the fact that pollen grains develop inside a cavity surrounded by different cell layers having various functions. Some anther cells can be partly autonomous from a nutritive point of view--that is, photosynthetic--but this is not a general rule. The process of development seems to have some resting sites, in the sense that sometimes, in the cells of the anther wall, in the loculus, or inside the microspore or vegetative cell, there are storage products, mainly starch and lipids. This temporary storage can be due to the fact that the anther represents a sinking site only for part of its development, namely, until the first haploid mitosis {Clement et al., 1996a). After that stage, all of the storage substances are consumed or translocated, and the anther wall cells are almost devoid of a cytoplasmic content when pollen grains are shed.

Why such different pathways in anther-plastid differentiation? One reason may be pollination type, which influences the resorption (or not) of tapetal elaioplasts and the physicochemical properties of pollenkitt. Whether or not the flower or the anther is photosynthetical may be another reason. A third may be whether the flower is produced before, at the same time as, or after the leaves. In the first case either the flower and the anther develop with substances accumulated earlier whereas in the latter two cases they can develop with freshly photosynthesized materials or both. Fourth, how long the process of pollen development lasts implies resting periods and deposits of substances. Finally, whether the plant lives in a dry environment or a wet one may influence the path of differentiation. Now we know that pollen grains with carbohydrate localized in the cytoplasm rather than in the plastids survive better and longer than do those with starch grains (Speranza et al., 1997).

Plastid differentiation in the vegetative cell is commonly a one-way process; in the reproductive cells, however, it seems different because there are some cycles of differentiation/dedifferentiation as proplastid/amyloplast/proplastid.

The description of plastid differentiation and dedifferentiation occurs in fixed, dehydrated, and embedded material observed mainly with electron microscopy. This technique is used because of the higher level of resolution but has negative aspects, such as it shows cells in rigor mortis. Certainly, plastids move inside the cell, especially around vacuoles, and a few studies suggest plastid movement. Pacini and Juniper (1979b) suggested that tapetal cell plastids, before they become elaioplasts, move around the vacuole. The movement of cytoplasm and plastids is probably involved in the secretion of tapetal cells. Ripe pollen is, with a few exceptions, devoid of vacuoles because it is usually dormant. But even in this case some exceptions have been noted--for example, in grasses such as Triticum aestivum, in which pollen amyloplasts move at shedding and during dispersal because, in grasses, pollen grains do not have a period of dormancy (Heslop-Harrison et al., 1997).

Anther cell layers are organized like Russian dolls around the pollen locule, and, despite their closeness, they play very different functions (Fig. 7), which are normally devoted to entire plant tissues in vegetative organs. Such an organization means that cell differentiation and physiology are accurately controlled during anther ontogenesis and then pollen development. Furthermore, the variety of plastids and related functions in this organ is an illustration of this regulation process.

XI. Literature Cited

Bellani, L. M., E. Pacini & G. G. Franchi. 1985a. In vitro pollen grain germination and starch content in species with different reproductive cycle, I. Lycopersicum peruvianum Mill. Acta Bot. Neerl. 34: 59-64.

-----, ----- & -----. 1985b. In vitro pollen grain germination and starch content in species with different reproductive cycle, II. Malus domestica Borkh. Cultivars Starkrimson and golden delicious. Acta Bot. Neerl. 34: 65-71.

Bhadula, S. K. & V. K. Sawhney. 1989. Amylolytic activity and carbohydrate levels during the stamen ontogeny of a male fertile, and a "gibberellin-sensitive" male sterile mutant of tomato (Lycopersicon esculentum). J. Exp. Bot. 40: 789-794.

Biddle, J. A. 1979. Anther and pollen development in garden pea and cultivated lentil. Canad. J. Bot. 57: 1883-1900.

Bird, J., E. K. Porter & H. G. Dickinson. 1983. Events in the cytoplasm during male meiosis in Lilium. J. Cell Sci. 59: 27-42.

Caredda, S. & C. Clement. 1999. Androgenesis and albinism in Poaceae: Influence of genotype and carbohydrates. Pp. 211-228 in C. Clement, E. Pacini & J. C. Audran (eds.), Anther and pollen: From biology to biotechnology. Springer-Verlag, Berlin.

-----, P. Devaux, R. S. Sangwan & C. Clement. 1999. Differential development of plastid during microspore embryogenesis in barley. Protoplasma 208: 248-256.

-----, C. Doncoeur, P. Devaux, R. S. Sangwan & C. Clement 2000. Plastid differentiation during pollen development and microspore embryogenesis in Hordeum vulgare L. Sexual Pl. Reprod. 13: 95-104.

Christensen, J. E. & H. T. Horner Jr. 1974. Pollen pore development and its spatial orientation during microsporogenesis in the grass Sorghum bicolor. Amer. J. Bot. 61: 604-623.

Ciampolini, F., M. Nepi & E. Pacini. 1993. Tapetum development in Cucurbita pepo (Cucurbitaceae). Pp. 13-22 in M. Hesse, E. Pacini & M. T. M. Willemse (eds.), The tapetum: Cytology, function, biochemistry and evolution. PI. Syst. & Evol., Suppl. 7. Springer-Verlag, Vienna.

Clement, C. & J. C. Audran. 1995. Anther wall layers control pollen sugar nutrition in Lilium. Protoplasma 187: 172-181.

----- & -----. 1999. Anther carbohydrates during in viva and in vitro pollen development. Pp. 71-90 in C. Clement, E. Pacini & J. C. Audran (eds.), Anther and pollen: From biology to biotechnology. Springer-Verlag, Berlin.

-----, L. Chavant, M. Burrus & J. C. Audran. 1994. Anther starch variations in Lilium during pollen development. Sexual Pl. Reprod. 7: 347-356.

-----, M. Burrus & J. C. Audran. 1996a. Floral organ growth and carbohydrate content during pollen development in Lilium. Amer. J. Bot. 83: 459-469.

-----, P. Mischler, M. Burrus & J. C. Audran. 1997. Characteristics of the photosynthetic apparatus and [CO.sub.2]-fixation in the flower bud of Lilium, 2. Anther. Int. J. Pl. Sci. 158: 801-810.

-----, P. Laporte & J. C. Audran. 1998. The loculus content and tapetum during pollen development. Sexual Pl. Reprod. 11:94-106.

Corriveau, J. L., L. J. Goff & A. W. Coleman. 1990. Plastid DNA is not detectable in the male gametes and pollen tubes of an angiosperm (Antirrhinum majus) that is maternal for plastid inheritance. Curr. Genet. 17: 439-444.

Day, A. & T. H. N. Ellis. 1984. Chloroplast DNA deletions associated with wheat plants regenerated from pollen: Possible basis for maternal inheritance of chloroplasts. Cell 39: 359-368.

----- & -----. 1985. Deleted forms of plastid DNA in albino plants from cereal anther culture. Curr. Genet. 9: 671-676.

Dickinson, H. G. 1973. The role of plastids in the formation of pollen grain coatings. Cytobios 8:25-40.

-----. 1981. The structure and chemistry of plastid differentiation during male meiosis in Lilium henryi. J. Cell Sci. 52: 223-241.

----- & D. Lewis. 1973. The formation of tryphine coating the pollen grains of Raphanus and its properties relating to the self incompatibility system. Proc. Roy. Soc. London, B, 184: 149-156.

Ellis, T. H. N. & A. Day. 1986. A hairpin plastid genome in barley. EMBO J. 5: 2769-2774.

Feijo, J. A. & M. S. S. Pais. 1988. Ultrastructural modifications of plastids and starch metabolism during the microsporogenesis of Ophrys lutea (Orchidaceae). Ann. Bot. 61: 215-219.

Fitzgerald, M. A., S. H. Barnes, S. Blackmore, D. M. Calder & R. B. Knox. 1994. Pollen development and cohesion in a mealy and a hard type of orchid pollinium. Int. J. Pl. Sci. 155: 481-491.

Franchi, G. G. & E. Pacini. 1988. Pollen polysaccharide reserves in some plants of economic interest. Pp. 90-91 in M. Cresti, P. Gori & E. Pacini (eds.), Sexual reproduction in higher plants: Proceedings of the Tenth International Symposium on the Sexual Reproduction in Higher Plants, 30 May- 4 June 1988, University of Siena, Siena, Italy. Springer-Verlag, Berlin.

-----, ----- & P. Rottoli. 1984. Pollen viability in Parietaria judaica L. during the long blooming period and correlation with meteorological conditions and allergic diseases. Giorn. Bot. Ital. 118: 163-178.

-----, L. Bellani, M. Nepi & E. Pacini. 1996. Types of carbohydrates reserves in pollen: Localization, systematic distribution and ecophysiological significance. Flora 191:143-159.

Garrido, D., O. Vicente, E. Heberle-Bors & M. I. Rodriguez-Garcia. 1995. Cellular changes during the acquisition of embryogenic potential in isolated pollen grains of Nicotiana tabacum. Protoplasma 186:220-230.

Gori, P. 1982. Accumulation of polysaccharides in the anther cavity of Allium sativum, clone Piemonte. J. Ultrastruct. Res. 81: 158-162.

Hageman, R. & M. B. Schroder. 1989. The cytological basis of the plastid inheritance in angiosperms. Protoplasma 152: 57-64.

Hause, G. 1991. Ultrastructural investigations of mature embryo sacs of Daucus carota, D. aureus, and D. muricatus: Possible cytological explanations of paternal plastid inheritance. Sexual Pl. Reprod. 4: 288-292.

Heslop-Harrison, J. 1968. Tapetal origin of pollen coat substances in Lilium. New Phytol. 67:779-786.

-----, Y. Heslop-Harrison & J. S. Heslop-Harrison. 1997. Motility in ungerminated grass pollen: Association of myosin with polysaceharide-containing wall-precursor bodies (P-particles). Sexual Pl. Reprod. 10: 65-66.

Hess, M. 1991. Ultrastructure of organelles during microsporogenesis in Tillandsia pallidoflavens (Bromeliaceae). Pl. Syst. & Evol. 176: 63-74.

----- & M. Hesse. 1994. Ultrastructural observations on anther tapetum development of freeze-fixed Ledebouria socialis Roth (Hyacinthaceae). Planta 192: 421-430.

Hesse, M. 1993. Pollenkitt development and composition in Tilia platyphyllos (Tiliaceae) analysed by conventional and energy filtering TEM. Pp. 39-52 in M. Hesse, E. Pacini & M. T. M. Willemse (eds.), The tapetum: Cytology, function, biochemistry and evolution. Pl. Syst. & Evol., Suppl. 7. Springer-Verlag, Vienna.

----- & M. Hess. 1993. Recent trends in tapetum research: A cytological and methodological review. Pp. 127-145 in M. Hesse, E. Pacini & M. T. M. Willemse (eds.), The tapetum: Cytology, function, biochemistry and evolution. Pl. Syst. & Evol., Suppl. 7. Springer-Verlag, Vienna.

Hixon, R. M. & B. Brimhall. 1968. Waxy cereals and red iodine starches: Starch and its derivatives. Ed. 4. Chapman & Hall, London.

Hourcade, D. E., M. Bugg & D. F. Loussaert. 1986. The use of Gaspe variety for the study of pollen and anther development in maize. Pp. 319-324 in D. L. Mulcahy, G. B. Mulcahy & E. Ottaviano (eds.), Biotechnology and ecology of pollen: Proceedings of the International Conference on the Biotechnology and Ecology of Pollen, 9-11 July 1985, University of Massachusetts, Amherst, MA, U.S.A. Springer-Verlag, New York.

Jahne, A. & H. Lorz. 1995. Cereal microspore culture. Pl. Sci. (Elsevier) 109: 1-12.

Keijzer, C. J. 1987. The process of anther dehiscence and pollen dispersal, 1. The opening mechanism of longitudinally dehiscing anthers. New Phytol. 105: 487-498.

----- & M. T. M. Willemse. 1988a. Tissue interactions in the developing locule of Gasteria verrucosa during microsporogenesis. Acta Bot. Neerl. 37: 475-492.

----- & -----. 1988b. Tissue interactions in the developing locule of Gasteria verrucosa during microgametogenesis. Acta Bot. Neerl. 37: 493-508.

Kirichenko, A. B., E. B. Kirichenko & A. A. Chebotar. 1977. Ultrastructure of anther of Hordeum vulgare L. at the stage of bicellular pollen: Characteristics of plastid differentiation. Physiol Rast, 24: 751-755.

Kirichenko, E., T. Krendeleva, G. Koukarskikh & N. Nizovskaia. 1992. Structure et activite fonctionnelle des chloroplastes des antheres et du pericarpe des caryopses de b1e et de seigle. Compt. Rend. Acad. Sci. Paris, Ser. 3, Sci. Vie 314: 365-370.

-----, -----, ----- & -----. 1993. Photochemical activity in chloroplasts of anthers and caryopsis in cereals. Russ. Pl. Physiol. 40: 229-233.

Knox, R. B. & C. Suphioglu. 1996. Environmental and molecular biology of pollen allergens. Trends P1. Sci. 1: 156-164.

Kott, L. S., L. Polsoni & W. D. Beversdorf. 1988. Cytological aspects of isolated microspore culture of Brassica napus. Canad. J. Bat. 66: 1658-1664.

Lalonde, S., D. U. Beebe & H. S. Saini. 1997. Early signs of wheat anther development associated with the induction of male sterility by meiotic-stage water deficit. Sexual Pl. Reprod. 10: 40-48.

Linskens, H. F., P. van der Werken & W. Jorde. 1980. The formation of allergens during development of rye pollen (Secale cereale). Allergol. & Immunopathol. 8: 35-41.

Lisci, M., C. Tanda & E. Pacini. 1994. Pollination ecophysiology of Mercurialis annual. (Euphorbiaceae): An anemophilous species flowering all year round. Ann. Bat. 74: 125-135.

Ljubesic, N., M. Wrischer & Z. Devide. 1991. Chromoplasts--The last stages in plastid development. Int. J. Dev. Biol. 35: 251-258.

Lord, E. M. 1981. Cleistogamy: A tool for the study of floral morphogenesis, function and evolution. Bot. Rev. (Lancaster) 47:421-449.

Mandaron, P., M. F. Niogret, R. Mache & F. Moneger. 1990. In vitro protein synthesis in isolated microspores of Zea mays at several stages of development. Theor. Appl. Genet. 80: 134-138.

Miyamura, S., T. Kuroiwa & T. Nagata. 1987. Disappearance of plastid and mitochondrial nucleoids during the formation of generative cells of higher plants revealed by fluorescence microscopy, Protoplasma 141:149-159.

Mlodzianowski, F. & K. Idzikowska. 1978. The ultrastructure of anther wall and pollen of Hordeum vulgare at the microspore stage. Acta Soc. Bot. Poloniae. 47: 219-224.

Mogensen, H. L. 1996. The hows and whys of cytoplasmic inheritance in seed plants. Amer. J. Bot. 83: 383-404.

Murgia, M., M. Charzynska, M. Rougier & M. Cresti. 1991. Secretory tapetum of Brassica oleracea L.: Polarity and ultrastructural features. Sexual Pl. Reprod. 4: 28-35.

Nagata, N., C. Saito, A. Sakai, H. Kuroiwa & T. Kuroiwa. 1999a. The selective increase or decrease of organellar DNA in generative cells just after pollen mitosis one controls cytoplasmic inheritance. Planta 209: 53-65.

-----, -----, -----, ----- & -----. 1999b. Decrease in mitochondrial DNA and concurrent increase in plastid DNA in generative cells of Pharbitis nil during pollen development. Eur. J. Cell Biol. 78: 241-248.

Nepi, M., F. Ciampolini & E. Pacini. 1996. Plastid differentiation during Cucurbita pepo (Cucurbitaceae) pollen grain development. Sexual Pl. Reprod. 9:17-24.

Noher de Halac, I., I. A. Cismondi & C. Harte. 1990. Pollen ontogenesis in Oenothera: Comparison of genotypically normal with the male-sterile mutant sterilis. Sexual Pl. Reprod. 3:41-53.

Olmedilla, A., J. A. M. Schrauwen & G. J. Wullems. 1991. Visualization of starch-sybthase expression by in situ hybridization during pollen development. Planta 184: 182-186.

Pacini, E. 1994. Cell biology of anther and pollen development. Pp. 289-308 in E. G. Williams, A. E. Clarke & R. B. Knox (eds.), Genetic control of self-incompatibility and reproductive development in flowering plants. Kluwer Academic, Dordrecht, Netherlands.

-----. 1996. Types and meaning of pollen carbohydrate reserves. Sexual Pl. Reprod. 9: 362-366.

-----. 1997. Tapetum character states: Analytical keys for tapetum types and activities. Canad. J. Bot. 75: 1448-1459.

----- & G. G. Franchi. 1983. Pollen grain development in Smilax aspera L. and possible functions of the loculus. Pp. 183-190 in D. L. Mulcahy & E. Ottaviano (eds.), Pollen: Biology and implications for plant breeding: Proceedings of the Symposium on Pollen--Biology and Implications for Plant Breeding, Villa Feltrinelli, Lake Garda, Italy, June 23-26, 1982. Elsevier Biomedical, New York.

----- & -----. 1991. Diversification and evolution of the tapetum. Pp. 301-316 in S. Blackmore & S. H. Barnes (eds.), Pollen and spores: Patterns of diversification. Systematics Association; Clarendon Press, Oxford.

----- & B. E. Juniper. 1979. The ultrastructure of pollen-grain development in the olive (Olea europea), 2. Secretion by the tapetal cells. New Phytol. 83: 165-174.

----- & -----. 1984. The ultrastructure of pollen grain development in Lycopersicum peruvianum. Caryologia 37: 21-50.

-----, P. E. Taylor, M. B. Singh & R. B. Knox. 1992a. Development of plastids in pollen and tapetum of rye-grass, Lolium perenne L. Ann. Bot. 70: 179-188.

-----, -----, -----, & -----. 1992b. Plastid developmental pathways in some angiosperm reproductive cells. Pp. 36-42 in E. Ottaviano, D. L. Mulcahy & M. Sari-Gorla (eds.), Angiosperm pollen and ovules. Springer-Verlag, New York.

Panchaksharappa, M. G. & C. K. Rudramuniyappa. 1974. Localization of nucleic acids and insoluble polysaccharides in the anther of Zea mays L.: A histochemical study. Cytology 39: 133-138.

Pandolfi, T., E. Pacini & D. M. Calder. 1993. Ontogenesis of monad pollen in Pterostylis plumosa (Orchidaceae, Neottiaideae). P1. Syst. & Evol. 186: 175-185.

Pifanelli, P., J. E. Ross & D. J. Murphy. 1998. Biogenesis and function of the lipidic structures of pollen grains. Sexual P1. Reprod. 11: 65-80.

Pyke, K. A. 1997. The genetic control of plastid division in higher plants. Amer. J. Bot. 84: 1017-1027.

Reznickova, A. 1978. Histochemical study of reserve nutrient substances in Lilium candidum. Compt. Rend. Acad. Bulgarie Sci. 31: 1067-1070.

----- & H. G. Dickinson. 1982. Ultrastructural aspects of storage lipid mobilization in the tapetum of Lilium hybrida var. Enchantment. Planta 155: 400-408.

----- & M. T. M. Willemse. 1980. Formation of the pollen in the anther of Lilium, 2. The function of the surrounding tissues in the formation of pollen and pollen wall. Acta Bot. Neerl. 29: 141-156.

----- & -----. 1981. The function of the tapetal tissue during microsporogenesis in Lilium. Acta Soc. Bot. Poloniae 50: 83-87.

Saini, H. S. 1997. Effects of water stress on male gametophyte development in plants. Sexual P1. Reprod. 10: 67-73.

----- & S. Lalonde. 1998. Injuries to reproductive development under water stress, and their consequences for crop productivity. J. Crop Prod. 1: 223-248.

Sangwan, R. S. & B. S. Sangwan-Norreel. 1987. Ultrastructural cytology of plastids in pollen grains of certain androgenic and nonandrogenic plants. Protoplasma 138: 11-22.

Schroder, M. B. 1985. Ultrastructural studies on plastids of generative cells in Liliaceae. 3. Plastid distribution during pollen development in Gasteria verrucosa (Mill.) Duval. Protoplasma 124: 123-129.

Schumann, C. M. & S. M. Hancock. 1989. Paternal inheritance of plastids in Medicago sativa. Theor. Appl. Genet. 78: 863-866,

Sodmergen, H., T. Suzuki, S. Kawano, S. Nakamura, S. Tano & T. Kuroiwa. 1992. Behavior of organelle nuclei (nucleoids) in generative and vegetative cells during maturation of pollen in Lilium 1ongiflorum and Pelargonium zonale. Protoplasma 168: 73-82.

-----, Y. Y. Luo, T. Kuroiwa & S. Y. Hu. 1994. Cytoplasmic DNA apportionment and plastid differentiation during male gametophyte development in Pelargonium zonale. Sexual P1. Reprod. 7: 51-56.

-----, H. Bai, J. X. He, H. Kuroiwa, S. Kawano & T. Kuroiwa. 1998. Potential for biparental cytoplasmic inheritance in Jasminum officinale and Jasminum nudiflorum. Sexual P1. Reprod. 65: 107-112.

Speranza, A., G. L. Calzoni & E. Pacini. 1997. Occurrence of mono- or disaccharides and polysaceharide reserves in mature pollen grains. Sexual P1. Reprod. 10:110-115.

Sun, C. S., S. C. Wu, C. C. Wang & C. C. Chu. 1979. The deficiency of soluble proteins and plastid ribosomal RNA in the albino plantlets of rice. Theor. Appl. Genet. 55: 193-197.

Sunderland, N. & B. Huang. 1985. Barley anther culture: The switch of programme and albinism. Hereditas (Lund), Suppl. 3: 27-40.

Takahashi, M. 1987. Development of omniaperturate pollen in Trillium kamtschaticum (Liliaceae). Amer. J. Bot. 74: 1842-1852.

Tanaka, I. 1991. Microtubule-determined plastids distribution during microsporogenesis in Lilium longiflorum. J. Cell Sci. 99: 21-31.

Taylor, P. E., K. Spuck, P. M. Smith, J. M. Sasse, T. Yokota, P. G. Griffiths & D. W. Cameron. 1993. Detection of brassinosteroids in pollen of Lolium perenne L. by immunocytochemistry. Planta 189: 91-100.

Ting, J. T. L., S. S. H. Wu, C. Ratnayake & A. H. C. Huang. 1998. Constituents of tapetasomes and elaioplasts in Brassica campestris tapetum and their degradation and retention during microsporogenesis. P1. J. 16 (5): 541-551.

Wang, M., S. Hoekstra, S. van Bergen, G. E. M. Lamers, B. J. Oppedjik, W. de Priester & R. A. Schilperoort. 1999a. Apoptosis in developing anthers and the role of ABA in this process during androgenesis in Hordeurn vulgare L. P1. Molec. Biol. 39: 489-501.

-----, S. van Bergen, G. E. M. Lamers, B. J. Oppedjik & R. A. Schilperoort. 1999b. Programmed cell death during androgenesis in Hordeurn vulgare L. Pp. 201-210 in C. Clement, E. Pacini & J. C. Audran (eds.), Anther and pollen: From biology to biotechnology. Springer-Verlag, Berlin.

Weber, M. 1992. The formation of pollenkitt in Apium nodiflorum (Apiaceae). Ann. Bot. 70: 573-577.

-----, 1996. The existence of a special exine coating in Geranium robertianum pollen. Int. J. P1. Sci. 157: 195-202.

Wetzel, C. L. & W. A. Jensen. 1992. Studies of pollen maturation in cotton: The storage reserve accumulation phase. Sexual P1. Reprod. 5: 117-127.

Wheatley J. M. 1977. Variations in the basic pathway of chloroplast development. New Phytol. 78: 407-420.

Xi, X. Y. 1991. Development and structure of pollen and embryo sac in Peanut (Arachis hypogaea L.). Bot. Gaz. (Crawfordsville) 152: 164-172.

Zaki, M. A. & H. G. Dickinson. 1990. Structural changes during the first divisions of embryos resulting from anther and free microspore culture in Brassica napus. Protoplasma 156: 149-162.

Zavada, M. S. 1984. Pollen wall development in Austrobaileya maculata. Bot. Gaz. (Crawfordsville) 145: 11-21.
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