Monday, 20 May 2019

Pollination and fertilization in plants

Pollination and fertilization in plants



This paper focuses on events post-pollination, and will only describe the forms of pollen, stigma, style, ovary, and embryo sac where the structure is relevant to the process being discussed. Data have been included on a selective basis; the paper, therefore, must not be regarded as an exhaustive analysis. For further information, a number of reviews have recently appeared relating to pollination and fertilization. These include reviews of the male gametophyte (Mascarenhas, 1989), pollen, pistil and reproductive function (Knox & Williams, 1986), pollen germination and tube growth (Heslop-Harrison, 1987; Steer & Steer, 1989), micro- and megasporogenesis and fertilization (Fougere-Rifot, 1987), the cytological basis of the plastid inheritance in angiosperms (Hagemann & Schroder, 1989), and molecular aspects of fertilization (Cornish, Anderson & Clarke, 1988). Self-incompatibility has also been deliberately given a low priority in this paper because excellent reviews have been published (Gibbs, 1986; Heslop-Harrison, 1983; Clarke et al., 1989) and in order to emphasize other aspects of the process of fertilization and its control. Where relevant in any discussion, self-incompatibility is briefly considered.

The pollen-stigma interaction
Stigma receptivity
The development of receptivity of the stigmatic surface may vary widely between species from several days prior to anthesis to several days after the flower opens (Palser, Rouse & Williams, 1989). In 'wet' stigmas receptivity often coincides with the accumulation of stigmatic secretion and in most stigmas with a positive response for non-specific esterase (HeslopHarrison and Shivanna, 1977). In protogynous and protandrous species stigma receptivity is strictly related to anther dehiscence (Richards, 1986). Some responses appear very specialized. In Talinum menagerie(Portulacaceae) (Murdy & Carter, 1987), for example, pollen germination is delayed for up to 2 h after compatible pollination. 

The delay depends entirely on the onset of stigma receptivity. Murdy & Carter (1987) consider that this trait promotes the accumulation of pollen on the stigma and simultaneous pollen germination, which in turn may intensify male gametophyte competition. It also may encourage pollen to be deposited from more than one plant on a stigma, a situation which has been shown to be positively correlated with seed mass per fruit (Marshall & Ellstrand, 1986) and the quality of sporophytic offspring (Van der Kloet, 1984), although such a correlation appears not to be universal (Schemske & Pautler, 1984; Bertin, 1986). There is no published case of pollen touch-down initiating stigma receptivity; it appears to be a preprogrammed response.

Adherence, hydration, recognition
A number of common events have been identified following pollen touchdown on the stigma incompatible pollination. Pollen adherence (particularly necessary for 'dry' stigma forms), pollen hydration (more accurately rehydration, since pollen grains, lose water prior to anther dehiscence (Heslop-Harrison, 1987)) and pollen recognition (particularly in self-incompatible species) are characteristics that take place before pollen germination (Heslop-Harrison, 1987). There is some circumstantial evidence to indicate a sequential process (pollen adherence, for example, may be a prerequisite for pollen rehydration) and for the stigma to possess the machinery for ensuring a carefully regulated rehydration of pollen grains before their germination (Heslop-Harrison, 1979a,b).

Pollen germination
Pollen germination is temperature-dependent. An extreme example is found in Juglans regia or Juglans nigra cultivars where no pollen germinates on stigmas below 14 °C or at 40 °C (Luza, Polito & Weinbaum, 1987). Pollen germination rates are, however, very variable between species (Table 1). Grass pollen appears to germinate most rapidly in vivo and pollen of Orchidaceae most slowly. Although a considerable body of information is available on stigmatic surface secretions (Knox, 1984; Dumas et al, 1988) there is no adequate explanation for such variation.

There are two possible candidates, however. 'Dry' stigmas appear to be associated with more rapid germination, although data are very limited, and trinucleate grains appear to germinate more rapidly than binucleate (Hoekstra, 1979; Hoekstra & Bruinsma, 1979). The grasses are surely a special case: their pollens are often very short-lived (measured in minutes rather than hours) and their stigmas remain receptive for short periods, perhaps not more than 3 ^ h (Heslop-Harrison, 19796). Using anhydrous fixation techniques, Dickinson & Elleman (1985) have shown that the pollen grain coat and the stigma papillar surface appear adapted for strict control of water entry into the grain. In addition, different self-incompatibility (S) genotypes have been shown to have significant differences in stigma cuticle thickness, which may itself influence the rate of pollen hydration (Elleman et ai, 1988).

It should not be surprising that in both self-incompatible and self-compatible species, not all pollen germinates on the stigma or that some tubes abort on the stigma (Palser et ai, 1989) or in the style. Both the genotype of the pollen and the environment may affect pollen viability and preclude germination or reduce and slow tube growth (Thomson, 1989).

Stylar penetration
Pollen germination is followed by pollen tube growth. In species with 'dry' stigma forms (e.g. Crocus (Heslop-Harrison & Heslop-Harrison, 1975) or Brassica oleracea (Roberts et ai, 1980; Elleman et al, 1988)), penetration of the stigmatic cuticle is necessary before the pollen tubes can enter the style and grow towards the ovary, whereas in many other species the cuticle is ruptured during the normal development of the receptive stigma (Dickinson & Lawson, 1975) or during the act of insect visitation and pollination (e.g. Papilionoideae-Leguminosae) (Lord & Heslop-Harrison, 1984; Kreitner & Sorensen, 1984; Owens, 1985).

Post-pollination stigmatic responses
The stigma in Acacia species is cup-shaped and contains a drop of secretion, which is considered to aid pollen adhesion (Kenrick & Knox, 1981). Pollination, however, promotes the development of a massive post-pollination exudate within approximately 30 min of touch-down, regardless of the pollen's origin. Kenrick & Knox (1981) postulate that this is the result of a general pollination signal, which is initiated at the commencement of the pollen-stigma interaction. Post-pollination responses are readily observable in the flowers of Orchidaceae. Stigma closure 24 h after either a self- or cross-pollination and column and ovary swelling, are characteristic for the many members of the family (Clifford & Owens, 1988). There is considerable evidence to suggest that such responses are promoted by auxin; auxin has very recently been found in pollinia of orchids by using thin-layer chromatography techniques (S.C. Clifford & S.J. Owens, unpublished results). Pollination also shortens the time to the onset of a flower wilting, petal color change, and abscission (Arditti, 1979; Lovell, Lovell & Nichols, 1987).

Pollen tube growth
Pre-programmed development of the style
Progressive, programmed development of the stylar transmitting tissue, which is known to be independent of pollination, has been found even though the activation of the pistil, as a consequence of pollination, is a well-known fact (Mulcahy & Mulcahy, 1988) and is dealt with later in this paper. Dickinson, Moriarty & Lawson (1982), for example, found that after the start of the stigmatic activity (papillar cell elongation, cuticle rupture, and secretion development) and before pollination, stylar canal cells developed in a 'wave' starting at the top of the pistil and continuing down the style. The stylar canal-cell cytoplasm became stratified, with nuclei, plastids and lipid droplets accumulating at sites away from the wall lining the canal whereas ER, dictyosomes and some mitochondria moved towards the canal wall. The canal lining wall itself developed transfer cell-like features. Recently, S.J. Owens (1989 and unpublished data) has shown that the breakdown of the intercellular connections of cells surrounding the stylar canal of Caesalpinia pulcherrima is part of a preprogrammed, post-anthesis development and are not stimulated by pollination.

Pollen tube extension
The pollen tube extends by tip growth (Steer & Steer, 1989); pollen tube growth may be through a stylar canal, intercellularly through a core of transmitting tissue or through an area of cellular debris, which appears more akin to growth in a stylar canal. In Liliaceae (Dickinson et al., 1982; Willemse & Franssen-Verheijen, 1988) and Leguminosae (Heslop-Harrison, 1982), pollen tube growth initially is through solid or semi-solid transmitting tissue, and subsequently through a stylar canal. The pollen tube has recently been the subject of major reviews (Heslop-Harrison, 1987; Steer & Steer, 1989), which have examined the current data on the tube cytoplasm, wall structure and the interaction of pollen tubes with calcium ions. Pollen tubes have a marked cytoplasmic polarity; distinct zones can be recognized in the cytoplasm based on their complements of nuclei and cells and cytoplasmic organelles. Dictyosomes are particularly important in tip growth: the vesicles supply both wall material and plasma membrane components (Steer & Steer, 1989). 

The tube wall comprises an outer pectic layer and an inner callosic-cellulosic layer, which is absent from the tube tip. The highest concentrations of calcium (both membrane-bound and free) are found at the tube tip. Some of the most interesting data concern the cytoskeleton, which has been examined by the use of fluorescent-labeled antibody techniques (Derksen, Pierson & Traas, 1985) and freeze-substitution (Lancelle, Cresti & Hepler, 1987). Pollen tube microtubules are arranged in axial or S-helical arrays (Derksen et al., 1985; Pierson, Derksen & Traas, 1986) and may be interspersed with and cross-linked to microfilaments (Lancelle et al., 1987) forming a structurally integrated cytoskeletal complex with the plasma membrane (Tiwari & Polito, 1988). Numerous bundles of microfilaments are found in the tube cytoplasm, particularly a three-dimensional network of short microfilaments in the tip region (Tiwari & Polito, 1988). Their functions involve wall organization, the movement of the vegetative nucleus and the generative cell or sperm cells, vesicle movement, and movement of cell organelles during cytoplasmic streaming (Heslop-Harrison et al., 1988). Myosin has recently been detected by immunofluorescence techniques on individual organelles in Alopecurus
pratensis and Secale cereale and on the surfaces of vegetative nuclei and generative cells in Hyacinthus orientalis and Helleborus foetidus (HeslopHarrison & Heslop-Harrison, 1989). These data indicate that the movement of cytoplasmic components in the pollen tube is driven by the interaction of myosin and actin fibrils at zones of contact (Heslop-Harrison & Heslop-Harrison, 1989). Contacts between the vegetative nucleus and the generative cell or sperm cells (Dumas et al., 1988) may also increase the efficiency and speed of movement.

Pollen tube growth in the style is believed to be heterotrophic and follows the path of the available reserves through the transmitting tissue. Starch digestion is triggered by pollination and only occurs incompatible matings in Petunia (Herrero & Dickinson, 1979). The time taken for the pollen tubes to reach the ovary varies greatly between species; this variation presumably indicates differences in style length, the form of the stylar transmitting tissue and ambient temperature. Heslop-Harrison (1987) has also suggested that the speed of wall synthesis is an important factor. Pollen tube growth rates are variable from 3 \xm s" 1 in maize and 2 \xm s" 1 in the rye (Heslop-Harrison 19796) to 0.3 - 1.4 [xm s" 1 in Lilium longiflorum (Van der Woude & Morre, 1967), Talinum mengesii (DuBay, 1981) and Gasteria verrucosa (Willemse & Franssen-Verheijen, 1988) to 0.07 - 0.14 pirn s" 1 in Crocus spp. (Heslop-Harrison, 1977) and Rhododendron nuttallii (Palser et ai, 1989).

 In all cases, however, the process appears to be dependent on the prevailing environmental conditions (Palser et al., 1989). A number of publications suggest that pollen tube growth is biphasic (Dickinson & Lawson, 1975) or triphasic (Willemse & FranssenVerheijen, 1988). Initial growth, particularly penetration of the stigma and growth in the neck of the style, is slow, but rapidly increases in growth rate to a maximum from this point. Recently, Willemse & Franssen-Verheijen (1988) have shown that initial growth in Gasteria verrucosa is slow (0.3 |im s" 1 ). Growth through the style is more rapid (1.4 \im s" 1 ); growth in the ovary returns to a rate similar to the initial growth rate (0.3 \im s" 1 ). Such changes in growth rate must reflect either limitation in nutritional resource or rapid metabolic change induced by the changing environments. The time taken for pollen tube growth from stigma to ovary varies from 30 to 60 min in Crepis capillaris (Gerassimova, 1933; Kuroiwa, 1989), Triticum aestivum (Bennett et al., 1973) and Hordeum vulgare (Engell, 1988) to 4-5 h in Gasteria verrucosa (Willemse & Franssen-Verheijen, 1988), 16 h in Salpiglossis sinuata (Hepher & Boulter, 1987), 60 h in Juglans regia (Luza et al., 1987), 8-9 din Rhododendron nuttallii (Palser et al., 1989) and 12 d in Prunus persica (Herrero & Arbeloa, 1989). In only the last species is the pollen tube seen to stop growth while the obturator becomes receptive to pollen tube passage (Herrero & Arbeloa, 1989).

Pollen tube guidance
The growth of the pollen tube in stigma, style, and ovary requires nutritional support; the structures of each organ or tissue may serve to orientate the pollen tube (Hepher & Boulter, 1987). Webb & Williams (1988) suggest that the guidance system for pollen tube growth through the pistol is likely to combine at least three factors. Firstly, pollen tubes probably follow paths of least physical resistance and tend to grow along surfaces because of their thigmotropic characteristic (Hirouchi & Suda, 1975). Gasser et al. (1989) isolated several pistil-specific cDNAs, one of which is expressed at anthesis but only in tissues subtending the stigma and in the outer layers of the transmitting tissue. 

They hypothesize that the expressed gene product is involved in preventing pollen tubes from leaving the transmitting tissue during pollen tube growth. Secondly, the stylar transmitting tract is assumed to provide the appropriate physical medium, nutritional factors (Labarca & Loewus, 1972, 1973), and recognition factors. Thirdly, chemotropism may impose directional tube growth (Miki-Hiroshige, 1964; Welk, Millington & Rosen, 1965; Mascarenhas, 1975; Picton & Steer, 1983; Mulcahy & Mulcahy, 1986) although evidence for this is not convincing (Heslop-Harrison, 1986) except, perhaps, in the ovary (Webb & Williams, 1988). Calcium has been identified as acting as a possible chemotropic agent (Steer & Steer, 1989). In experiments designed to test the effects of pollination on stylar changes, Mulcahy & Mulcahy (1988) pollinated Nicotiana alata flowers in mid-style and observed pollen tube growth. In unpollinated stigmas, pollen tubes grew both towards the stigmatic end and towards the ovary, suggesting no chemotropic gradient, whereas, in pollinations performed after astigmatic one, pollen tubes grew predominantly towards the ovary, suggesting that a gradient had been set up. 

Pollen tube growth in the ovary and entry to the
micropyle
Entry into the ovule from the transmitting zone is followed by tube growth over the ovarian transmitting zone (placental epithelium), which is intimately associated with the micropyle of the ovule and facilitates pollen tube contact. The transmitting zone may (Cass et ah, 1986; Clifford & Owens, 1990) or may not (Olson, 1988) be associated with placental exudate. The placentae of Papaver nudicaule produce an exudate through which pollen tubes must grow on their way to the ovular micropyles (Cass et ah, 1986); the exudate may be directly involved in the recognition of the compatible tube. The placentae of Solarium tuberosum are stimulated by successful pollination; placental cell proliferation and enlargement ensue (Olson, 1988).

Herrero, Arbeloa & Gascon (1988) reported that the period from arrival of pollen tubes at the base of the style to fertilization was variable in the peach (Prunus persica) and that this period could extend beyond the time taken for pollen-tube growth down the style. The principal cause of the delay appears to be the obturator, a placental protuberance connecting the style with the micropyle of the ovule, which at this time appears unreceptive to pollen tubes. Starch digestion takes place in the obturator and is concomitant with the secretion of proteins and carbohydrates; however, this development is not stimulated by pollination but appears to be a maturation process, since it occurs in both pollinated and unpollinated pistils (Arbeloa & Herrero, 1987). Hence pollen tubes must wait for its receptivity. Once pollen tubes have passed the obturator, callose begins to be deposited in this structure (Arbeloa & Herrero, 1987). This latter paper is the only demonstration of such a phenomenon, other workers observing pollen tube growth on the obturator but not observing such changes (Tilton & Horner, 1980; Tilton et al., 1984; Hill & Lord, 1987). Once the pollen tubes have traversed the obturator, there are further periods before micropylar entry and fertilization (Herrero et al., 1988). A chemotrophic effect may be necessary for penetration of the ovule (Lord & Kohorn, 1986). 

Fertilization
Pollen tube entry into the embryo sac
Generally, pollen tube entry into the micropyle is followed by intercellular growth through the nucellus to the embryo sac. Various modifications and elaborations of the micropylar area exist, such as the extension of the inner integument in Crocus to form a micropylar canal (Rudall, Owens & Kenton, 1984). The most bizarre cases, however, are where the micropylar ends of the synergid cells may be extended into the micropyle (and even up the style sometimes to the stigma in Viscaceae; Mahesh- wari, 1950) or modified into hooks. The embryo sac cell first encountered is generally one of the synergies. The cell wall at the micropylar end is often elaborated into the filiform apparatus, a labyrinthine wall modification of structure similar to that in transfer cells. Such a structure has led some authors to consider that these cells are necessarily involved in secretion and absorption (van Went, 1970; Chao, 1971; Wilms, 1981). A filiform apparatus is absent in Crepis capillaris (Kuroiwa, 1989).

Recently, from a study of 35000 ovules by using ovule-clearing techniques in Rhododendron nuttallii, it has become apparent that the ovule and its enclosed gametophyte are far from mature during the period of anthesis and pollination (Palser et aL, 1989). The earliest stage observed at anthesis was a linear tetrad; the majority of megagametophytes were at a stage earlier than four nucleate. In fact, no megagametophyte was mature at pollination. Some of the variations may be due to ovule position on the placenta (Palser et aL, 1989).

The order of pollen tube entries into ovules and subsequent fertilization may be related to variations in the rate of gametophyte maturation in different ovules as well as variations in the rate of pollen tube growth. Fertilized ovules constituted less than 5% of the total ovules until 12 d after pollination (Palser et aL, 1989). A number of investigations have indicated that pollination stimulates the degeneration of one of the two synergids (Table 2) and that pollen tube entry and sperm discharge in the embryo sac must occur via the degenerate synergid. The full story is yet to be revealed for variations in the structure and composition of the embryo sac are well known. The embryo sac may have one or no synergids (Plumbago capensis, Cass, 1972; P. zeylanica, Russell, 1982), and ephemeral through to multiple antipodal cells (e.g. Gramineae).

Penetration by more than one pollen tube is occasionally observed (Palser et aL, 1989). This has also been observed where wide crosses have been attempted between wheat and maize (Laurie & Bennett, 1988).

Nuclear fusion
Very little is known about the fusion of the nuclei of egg and sperm and polar nuclei and sperm. The fusion of the polar nuclei in the central cell may occur before fusion with the sperm nucleus or fusion may be simultaneous. At fertilization, the polar nuclei are still separate in Rhododendron nuttallii (Palser et al., 1989). The sperm nucleus becomes increasingly diffuse before it fuses with the egg nucleus in Triticum aestivum (Bennett et al, 1973). No sperm nucleus could be recognized between 18 and 20 h after pollination. In the central cell, the polar nuclei and sperm nucleus remained distinct at 2-3 h after pollination and still had separate nuclear membranes. At prophase of the first division, the sperm chromosomes remained distinct but increasingly merged with the polar nuclei as contraction progressed. The division was apparent at 5 h (Bennett et al., 1973). The observations of Bennett et al. (1973) differed from those of Morrison (1955), who reported that chromosomes were not visible as separate groups at the primary division of the endosperm nucleus. In Triticum aestivum, the first division of the zygote is seen at about 22 h from pollination; by 24 h after pollination, the endosperm consists of about 16 nuclei (Bennett et al., 1973). In Hordeum vulgare (Engell,\ 1988), sperm and egg nuclei were fused by 45 min after pollination, the first zygotic division being timed similarly to wheat at 22-24 h after pollination. Following the development of the single primary endosperm nucleus, a transitory callose wall is formed around the zygote in Rhododendron and Ledum species (Williams et al., 1984; Palser et al., 1989).

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