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. 2009 Nov 19;105(2):221–231. doi: 10.1093/aob/mcp276

The progamic phase of an early-divergent angiosperm, Annona cherimola (Annonaceae)

J Lora 1, J I Hormaza 1,*, M Herrero 2
PMCID: PMC2814751  PMID: 19939980

Abstract

Background and Aims

Recent studies of reproductive biology in ancient angiosperm lineages are beginning to shed light on the early evolution of flowering plants, but comparative studies are restricted by fragmented and meagre species representation in these angiosperm clades. In the present study, the progamic phase, from pollination to fertilization, is characterized in Annona cherimola, which is a member of the Annonaceae, the largest extant family among early-divergent angiosperms. Beside interest due to its phylogenetic position, this species is also an ancient crop with a clear niche for expansion in subtropical climates.

Methods

The kinetics of the reproductive process was established following controlled pollinations and sequential fixation. Gynoecium anatomy, pollen tube pathway, embryo sac and early post-fertilization events were characterized histochemically.

Key Results

A plesiomorphic gynoecium with a semi-open carpel shows a continuous secretory papillar surface along the carpel margins, which run from the stigma down to the obturator in the ovary. The pollen grains germinate in the stigma and compete in the stigma-style interface to reach the narrow secretory area that lines the margins of the semi-open stylar canal and is able to host just one to three pollen tubes. The embryo sac has eight nuclei and is well provisioned with large starch grains that are used during early cellular endosperm development.

Conclusions

A plesiomorphic simple gynoecium hosts a simple pollen–pistil interaction, based on a support–control system of pollen tube growth. Support is provided through basipetal secretory activity in the cells that line the pollen tube pathway. Spatial constraints, favouring pollen tube competition, are mediated by a dramatic reduction in the secretory surface available for pollen tube growth at the stigma–style interface. This extramural pollen tube competition contrasts with the intrastylar competition predominant in more recently derived lineages of angiosperms.

Key words: Annona cherimola, Annonaceae, embryo sac, endosperm, Magnoliid, ovule, pollen–pistil interaction, pollen tube

INTRODUCTION

Study of the reproductive biology of basal and early-divergent angiosperms has experienced a renaissance in recent years and is providing valuable information on evolutionary trends in flowering plants (Friedman and Ryerson, 2009; Rudall et al., 2009). Although much remains to be discovered about the basic features of the sexual process in angiosperms, a new understanding of its evolutionary developmental biology is beginning to emerge. Most angiosperms conform to a defined suite of reproductive characteristics but new data derived from studies on ancient extant flowering plant lineages reveal that the reproductive features of the first flowering plants differed significantly from those shown by the majority of extant flowering plants (Friedman and Williams, 2004). Recent work is revealing unique features in the female gametophyte of ancient extant angiosperms, such as the egg cell apparatus (Friedman, 2006), the endosperm (Friedman et al., 2008) and the provisioning of ovular resources (Friedman, 2008; Rudall et al., 2008). The study of pollen development and evolution is also emerging as a powerful field in understanding the evolution of reproductive characters (Rudall and Bateman, 2007; Lora et al., 2009a). Although information on the male and female side is accumulating in ancient lineages of extant angiosperms, there is still much to be learned about the interplay between the two: pollen–pistil interaction.

The progamic phase, the period of pollen tube growth through the pistil that elapses from pollination to fertilization and gamete fusion (Linskens, 1975; Williams, 2009), provides an opportunity for pollen–pistil interaction, which is emerging as a powerful strategy regulating mating in flowering plants (Herrero and Hormaza, 1996; Herrero, 2000, 2003; de Graaf et al., 2001; Rea and Nasrallah, 2008). The molecular mechanisms involved in this signalling are being deciphered (Escobar-Restrepo et al., 2007; Higashiyama and Hamamura, 2008; Hiscock and Allen, 2008), although a comprehensive view on the events and their implications remains to be made. Recent work in Amborella, sister to all extant angiosperms, sets the baseline for understanding the evolution of pollen–pistil interaction (Williams, 2009); in this sense, the study of the progamic phase in ancient angiosperms, which has been performed in a number of species belonging to ancient flowering plant lineages (Vithanage, 1984; Orban and Bouharmont, 1995; Pontieri and Sage, 1999; Thien et al., 2003; Sage and Sampson, 2003; Koehl et al., 2004; Hristova et al., 2005; Lyew et al., 2007), may prove a highly valuable tool to track the evolution of this process (Hiscock and Allen, 2008). Also, the evolution and function of the transmitting tissue in extant representatives of early-divergent angiosperm lineages has been recently explored (Sage et al., 2009).

However, one of the main limiting factors in studying developmental processes in early angiosperms and placing them in a phylogenetic framework is that most ancestral angiosperm lineages have arrived at the present time in a highly fragmented way. Early-divergent angiosperms have a very meagre representation, with some lineages represented by only one or very few species. Annonaceae is the largest living family in the early-divergent angiosperm clade magnoliids (APG II, 2003; Soltis et al., 2005), including about 130 genera and 2300 species with a worldwide distribution (Chatrou et al., 2004). Besides interest in the family regarding questions on early angiosperm evolution, some of the species in the Annonaceae, such as cherimoya (Annona cherimola), sugar apple (A. squamosa) or soursop (A. muricata), have been used as a food source by pre-Columbian cultures in South America (Popenoe, 1989), and they now have a clear niche for expansion in developing countries with subtropical climates. However, despite the importance for both basic and applied studies, very little is known of the reproductive biology of this family.

Most papers on the reproductive biology in the Annonaceae have focused on descriptive studies of the flowers (Norman et al., 1986, 1992; Carvalho and Weber, 2000; Norman, 2003; Kiill and Costa, 2003), embryology (Svoma, 1998a) and flower development (Decraene and Smets, 1990; Leins and Erbar, 1996). Studies on the pollen tube pathway are limited and restricted just to pollen germination in the stigma (Vithanage, 1984). Recently, pollen development and release in groups in A. cherimola have been studied (Lora et al., 2009b), and the coexistence of bi- and tricellular pollen at anther dehiscence in this species has been shown, contributing to the understanding of the heterochronic shift from bicellular to tricellular pollen (Lora et al., 2009a).

Here, the progamic phase and early embryo development is characterized in the early-divergent angiosperm A. cherimola. The examination of pistil anatomy reveals features showing a simple pollen–pistil interaction that are likely to be plesiomorphic for all angiosperms.

MATERIALS AND METHODS

Plant material

Annona cherimola shows protogynous dichogamy (Schroeder, 1971). Flowers are hermaphroditic, but female and male structures do not mature simultaneously, hindering self-fertilization in the same flower. Moreover, most flowers of the same genotype are synchronized and, consequently, transfer of pollen between different flowers of the same genotype is also difficult. The flower cycle is completed in 2 d; in the morning of the first day the flower is in preanthesis with the petals tightly closed. Around midday the flower passes to the female stage, petals slightly widen apart and the stigma is receptive. After approximately 30 h, flowers switch to the male stage. Anthers dehisce at approx. 1700–1800 h under the environmental conditions used herein. Concomitantly with anther dehiscence, the petals widen apart and the stigma dries up and loses receptivity.

Adult trees of A. cherimola ‘Campas’ located in a field cultivar collection at the EE La Mayora-CSIC, Málaga, Spain, were used in these experiments. Flowers were fixed along the flowering period, during two consecutive years.

Pollination procedures

Anthers and pollen were collected from flowers just before anther dehiscence, stored at 4 °C and used for hand pollination the following day. Fifteen flowers per day were pollinated at 0900 h on the first day of the flower cycle and the floral tube was then plugged with cotton to prevent further unwanted pollination. To evaluate developmental changes independent of pollination, similarly treated flowers were left unpollinated. The gynoecia of 15 pollinated and 15 unpollinated flowers were weighed and fixed daily from preanthesis (1 d before anthesis) to 3 weeks following hand pollination.

An additional group of flowers were collected in the field, placed in water in the laboratory at room temperature and pollinated at 0900 h on the first day of the flower cycle to study pollen tube growth. Regression analyses were used to describe the relationships between days after anthesis and pistil weight. Several regression models were tested and third-order polynomial regression was selected.

Microscopic preparations

Pollen tube growth was documented using squashed preparations of pistils from hand-pollinated flowers kept in water at room temperature. For this purpose, pistils were fixed in formalin–acetic acid–alcohol (FAA) 3, 6 and 9 h after pollination. Pistils were water washed and placed in 1 m NaOH for 1 h to soften the tissues. Individual pistils were dissected and squashed preparations were stained with 0·1 % aniline blue in PO4K3 (Currier, 1957; Linskens and Esser, 1957).

Following hand pollination in the field, pistils were also sequentially fixed. Eight flowers per day were collected at 0900 h and fixed in FAA, dehydrated in a graded ethanol series, and then embedded in paraffin wax. Seven flowers per day were fixed in 2·5 % glutaraldehyde in 0·03 m phosphate buffer (Sabatini et al., 1963). Finally, three flowers per day were fixed for 24 h in 3 : 1 (V1/V2) ethanol/acetic acid and transferred to 75 % ethanol for storage at 4 °C following the method of Williams et al. (1999). These flowers and those fixed in 2·5 % glutaraldehyde were dehydrated in a graded ethanol series and embedded in Technovit 7100 (Heraeus Kulzer, Wehrheim, Germany) resin.

To observe pollen tubes and callose, squashed preparations and paraffin-embedded material sectioned at 10 µm were stained with 0·1 % aniline blue in 0·1 n PO4K3 (Currier, 1957; Linskens and Esser, 1957). Sections were also stained with 0·01 % acridine orange in 0·03 m phosphate buffer to observe DNA and RNA (Nicholas et al., 1986; Dudley et al., 1987), with 0·01 % auramine O in 0·05 m phosphate buffer to observe cutine and suberine (Heslop-Harrison, 1977) and with 0·07 % calcofluor in water for cellulose (Hughes and McCully, 1975). For general histological examination, paraffin-embedded material was stained with a mixed staining in the following order: 0·1 % aniline blue in 0·1 n PO4K3, 0·01 % acridine orange in water, 0·01 % auramine O in water and 0·01 % calcofluor in water, for 10 min each at 40 °C to accelerate staining and drying. Others sections were also stained with safranin, crystal violet and green light according to Gerlach (1969). Resin-embedded material fixed in glutaraldehyde was sectioned at 2 µm and stained with periodic acid-Shiff's reagent (PAS) followed by 0·2 % toluidine blue in water (Feder and O'Brien, 1968) to observe insoluble carbohydrates and nuclei. Resin-embedded material in glutaraldehyde was also stained with iodine potassium iodide (IKI) for starch (Johansen, 1940). Resin-embedded material fixed with 3 : 1 (V1/V2) ethanol/acetic acid was sectioned at 5 µm and stained with a solution of 0·25 mg mL−1 4′,6-diamidino-2-phenylindole (DAPI) and 0·1 mg mL−1 p-phenylenediamine (added to reduce fading) in 0·05 m Tris (pH 7·2) for 1 h at room temperature in a light-free environment (Williams et al., 1999). Preparations were observed under an epifluorescence Leica DM LB2 microscope with 340–380-nm bandpass and 425-nm longpass filters for DAPI. Pollen tube growth rates were calculated as the length of the longest pollen tube divided by the time elapsed from pollination.

RESULTS

Gynoecium anatomy and pollen tube growth

The average number of carpels in the gynoecium of A. cherimola was 90·4 ± 5·4 (n = 5 flowers), and these were fused to form a syncarp occupying the centre of a conical receptacle. Each of the carpels had a single anatropous ovule that could develop into a single seed. The androecium was located below the gynoecium forming a helicoidal structure with up to 200 stamens. The pistil had a large stigma and a relatively short style with partial postgenital fusion at the periphery of the innermost side (Fig. 1A), forming an open stylar canal. The closing area of this canal was covered in a zip-like fashion by unicellular secretory papillae that resulted in a continuous secretory carpet starting in the stigma and going all the way through the style down to the ovary (Fig. 1B). Idioblastic solitary oil cells (Fig. 1C), thick-walled sclereid cells (Fig. 1D) and tanniferous cells were observed along the cortical tissue. Tanniferous cells were also observed in the ovular epidermis.

Fig. 1.

Fig. 1

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Gynoecium anatomy and pollen tube growth in Annona cherimola. (A) Pistil showing the stigma (stg), short style (stl) and ovary (ov) with partial postgenital fusion at the periphery of the innermost side (arrowhead). (B) Longitudinal section of the pistil showing pollen tube growth (arrowhead) through the short open stylar canal that leads to an anatropous ovule. (C) Oil cells. (D) Thick-walled sclereid cells. (E) Pollen tubes growing on the stigma towards the stigmatic furrow that leads to the stylar canal. (F) Stigma–style interface, with the stigmatic furrow leading to the narrow receptive closing margins of the stylar canal (arrowhead) and pollen tube growing through (pt). (G) Pollen tube (arrowhead) reaching the locule over the continuous papilar secretory zone. (H) Pollen tube growing through the micropyle formed by the inner integument (ii) that protrudes over a hood-shaped outer integument (oi), and reaching the nucellus (nu) 24 h after pollination. (A) Whole mount of a dissected pistil stained with aniline blue. (B, H) Aniline blue staining of a 10-μm paraffin section. (C, D) DAPI staining of a 5-μm resin section. (E, F) Aniline blue staining of squashed preparation. (G) Mixed staining of a 10-μm paraffin section. Scale bars: (A) 200 µm; (B) 200 µm; (C) 20 µm; (D) 20 µm; (E) 20 µm; (F) 100 µm; (G) 20 µm; (H) 20 µm.

The length of the pollen tube pathway from the stigma to the ovule was 1832·4 µm (±132·5, n = 10). Pollen grains germinated in the stigma and the pollen tubes grew on the stigmatic surface towards the suture line (Fig. 1E) to penetrate in the style. The stigmatic suture line formed a furrow along the entire stigma continuous with the style secretory area of the semi-open canal (Fig. 1F). Germination occurred rapidly and 1 h after pollination pollen grains had germinated and grew freely in the stigma. An average of 22·5 pollen grains (±15·2, n = 70 stigmas) were recorded per stigma with and average pollen germination of 43·6 % (±8·4 %, n = 70 stigmas). However, a drastic reduction in the number of pollen tubes occurred at the stigma–style interface (Fig. 1F), and only one to three pollen tubes were observed in the style. This reduction is related to the receptive surface available for pollen tube growth. Whereas the stigma was formed by a wide papillar surface 470·1 µm (±78, n = 10) in length and 307 µm (±69, n = 10) in width, the receptive surface in this area was restricted to the narrow stylar semi-closed margins that were lined by secretory papillae continuous with those of the stigma. This stigma–style interface was able to lodge very few pollen tubes which, in their way through the style, stick to this narrow receptive surface leaving the rest of the non-receptive stylar canal empty. Pollen tubes were first seen growing in the style 2–3 h after pollination, and travelled over 836·3 µm (±77·3, n = 10) to reach the ovary locule (Fig. 1G) some 4–6 h after pollination, with an average pollen tube growth rate in the style of 280 µm h−1. At the ovary locule, the pollen tubes grew over the obturator and penetrated the hood-shaped ovule after traversing a distance of 526 µm (±86, n = 10). A single pollen tube was observed penetrating each ovule (Fig. 1H) and the first fertilized ovules were observed 1 d after pollination.

Embryo sac and fertilization

Annona cherimola has an anatropous, bitegmic and crassinucellate ovule, with an endostomal micropyle formed by the inner integument that protrudes over the external integument (Fig. 2A). The outer integument is vascularized. At anthesis, the embryo sac is mature and shows the Polygonum-type structure with seven cells and eight nuclei, three at the micropylar end [the two synergids (Fig. 2A) and the egg cell (Fig. 2B)], three at the chalazal end [the three antipodal cells (Fig. 2C)] and two polar nuclei in the centre (Fig. 2D), which are not fused at anthesis.

Fig. 2.

Fig. 2

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Embryo sac in Annona cherimola. (A) Ovule showing the micropyle (asterisk) formed by the inner integument (ii), the shorter outer integument (oi) and the embryo sac with two synergids (sy). (B) Egg cell (ec). (C) Two of the three antipodal cells (arrowheads). (D) Two polar nuclei (pn). DAPI staining of 5-μm resin sections. Scale bars = 20 µm.

The embryo sac contains large starch grains distributed in the central cell (Fig. 3A). These starch grains are much larger than the standard starch grains located in the sporophytic tissues of the ovule. They react both to PAS and IKI stains. Starch accumulates around the egg cell (Fig. 3B), and also around the two polar nuclei (Fig. 3C) that fuse close to the time of fertilization. Both synergids have a large vacuole at the top of the cell (antipodal side) and the cytoplasm and the nucleus are located at the base (micropyle side) (Fig. 3D) where a filiform apparatus is developed (Fig. 3E). Starch disappears following fertilization concomitant with endosperm development. Endosperm is cellular and starts to develop 3 d after pollination. It has a bipolar nature where the first division produces a large cell in the micropylar pole and a smaller cell in the chalazal pole. Starch grains accumulate in the chalazal pole cell (Fig. 3F) and decrease with endosperm enlargement (Fig. 3G), disappearing 3 weeks after pollination concomitant with further endosperm cellular division. After fertilization, a zygote develops and the first cell division could be seen 8 d after pollination when the endosperm already has four cells (Fig. 3H).

Fig. 3.

Fig. 3

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Embryo sac and endosperm development in Annona cherimola. (A) Embryo sac of flower in preanthesis with starch grains (sg) around the two polar nuclei and showing the three antipodal cells (arrowhead). The difference between the standard starch grains (sg) located in the sporophytic tissues of the ovule and the big starch grains in the female gametophyte is apparent. (B) Egg cell with a large vacuole at the base of the cell. (C) Two polar nuclei surrounded by large starch grains. (D) Two synergid cells with nucleus and cytoplasm at the micropylar end and a large vacuole at the top of the cell. (E) Filiform apparatus (fa) of a synergid cell. (F) Starch grains (sg) accumulating in the chalazal pole of the embryo sac in a fertilized ovule four days after pollination. (G) Cellular endosperm 8 d after pollination with starch (sg) accumulated at the cell of the chalazal end. (H) Zygote first division 8 d after pollination. PAS (A, F, H) and PAS and toluidine blue (B–E, G) staining of a 2-μm resin section. Scale bars: (A) 20 µm; (B) 10 µm; (C) 10 µm; (D) 10 µm; (E) 10 µm; (F) 20 µm; (G) 20 µm; (H) 10 µm.

Changes in the pistil

The pollen tube pathway is lined by secretory papillae that form a continuous carpet from the stigma down to the placenta facing the ovule endostome. The papillae in the placenta resemble an obturator as they form a protuberance towards the ovule entrance. Cytohistologically, these papillae are similar to those of the stigma and style (Fig. 4A–C) although a basipetal maturation sequence can be seen from the stigma down to the obturator. The papillae in the less mature areas are rich in starch and show no secretion. As papillae mature, starch disappears concomitant with the production of a secretion on the surface of the papillae. Thus, although at preanthesis the stigma does not appear to contain starch (Fig. 4A), starch is still present in the style (Fig. 4B) where a secretion is being produced. At the obturator (Fig. 4C) starch is far more conspicuous and the secretion is not apparent at this time.

Fig. 4.

Fig. 4

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Secretion along the pollen tube pathway in Annona cherimola. Longitudinal section of secretory papillae of the (A) stigma, (B) style and (C) obturator in preanthesis with the same cytohistological features although with differences in maturation and starch content: while at the stigma (A) starch has already disappeared, in the style (B) starch is still present and a rich secretion is apparent; and the obturator (C) is still full of starch. (D) Transverse section of the style showing a semi-open stylar canal (arrowhead) lined only in the outermost side with papillar cells (pc) with secretion. Stylar longitudinal section in the outermost papillar secretory zone at preanthesis (E), showing starch grains. Same area, 1 d after pollination (F), shows less starch in the cells and secretion. Transverse section of obturator (ob) cells at preanthesis (G) with starch; (ii: inner integument) and 1 d after pollination (H) with little starch and abundant secretion (ii: inner integument; oi: outer integument). PAS and toluidine blue staining of 2-μm resin sections. Scale bars = 20 µm.

The secretion of the papillae is present along the whole pollen tube pathway from anthesis to 6 d later. This secretion stains heavily with PAS and with toluidine blue (Fig. 4B). The papillae and secretion are present just in the outermost side of the semi-open stylar canal (Fig. 4D). Before anthesis, the unicellular secretory papillae are rich in starch reserves in the style (Fig. 4B, E). One day after anthesis, starch disappears and the secretion increases (Fig. 4F) concomitant with pollen tube passage. The same situation can be seen in the obturator where starch is conspicuous before anthesis (Fig. 4G) but disappears as secretion increases 1 d later (Fig. 4H). This process does not seem to be triggered by the pollen tube passage, but appears to be developmentally regulated, as it occurs in the same way and at the same time in unpollinated flowers.

Although at preanthesis callose is not apparent in the papillae secretory cells, callose layering starts in the papillae 1 d after pollination (Fig. 5A) and is also present in the obturator (Fig. 5B). This callose layering occurs in a similar way in pollinated (Fig. 5B) and unpollinated flowers (Fig. 5C). However, callose layering in the nucellus at the base of the embryo sac appears 3 d after pollination only in pollinated flowers (Fig. 5D). By contrast, callose layering in the vascular bundles is only observed in ovules of unpollinated flowers 1 d after anthesis (Fig. 5E). During ovary development, callose also appears in the cell plates forming the walls of the cellular endosperm (Fig. 5F, G).

Fig. 5.

Fig. 5

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Callose layering along the pollen tube pathway and during early endosperm development in Annona cherimola. Callose 1 d after pollination in papillar cells (arrowhead) of style (A) and obturator (arrowhead) of a pollinated (B) and an unpollinated (C) flower. (D) Callose in the nucellus micropylar pole 3 d after anthesis. (E) Deposition of callose in vascular bundles of the ovule (arrowhead) of unpollinated flowers 1 d after pollination. Callose in the cell plates (arrowhead) forming the walls of the cellular endosperm, 4 d after pollination (F) and 8 d after pollination (G). Aniline blue staining of 10-μm paraffin sections. Scale bars = 20 µm.

Gynoecium weight increases slowly and is similar in pollinated and unpollinated flowers during the first 4 d after anthesis. Six days after anthesis differences can be observed among pollinated and unpollinated flowers; thus, whereas unpollinated flowers start to drop, pollinated flowers experience rapid growth (Fig. 6). A similar pattern was observed for pollinated and unpollinated flowers in the two years of observations.

Fig. 6.

Fig. 6

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Mean pistil weight after anthesis in pollinated and unpollinated flowers of Annona cherimola. Bars indicate s.d.

DISCUSSION

Annona cherimola shows a simple and plesiomorphic pistil, with a short style and a semi-open continuous secretory carpel, which supports a simple pollen–pistil interaction. Interestingly, this interaction exhibits a support–constrain strategy that is prevalent in phylogenetically derived angiosperm species (Herrero and Hormaza, 1996) although in the former case it takes place in the stigma rather than the style.

Pistil support to pollen tube growth

Annona cherimola has a wet stigma like other closely related species in this genus (Heslop-Harrison and Shivanna, 1977; Vithanage, 1984). In A. cherimola the common secretory papillar carpet that covers the stigma, style and ovary along the semi-suture line provides a substrate for pollen tube growth. Secretion along the pollen tube pathway has also been reported in other members of ancient angiosperm clades such as Trimenia moorei (Bernhardt et al., 2003), Illicium floridanum (Koehl, 2002, cited by Bernhardt et al., 2003), Amborella trichopoda (Thien et al., 2003), Saururus cernuus (Pontieri and Sage, 1999), Psedowintera axillaries (Sage and Sampson, 2003) and Kadsura longipedunculata (Lyew et al., 2007), and the substance involved appears to be composed of arabinogalactan and arabinogalactan-proteins (Sage et al., 2009). Although dry stigmas are considered as plesiomorphic in flowering plants (Thien et al., 2009) both dry and wet stigmas can be found in taxa of the ANITA and magnoliid clades (Thien et al., 2009). Molecular studies on wet and dry stigmas and their implications in pollen–pistil interaction have been performed only in a limited number of evolutionarily-derived angiosperm taxa and, consequently, there is a need for more studies on this topic among early-divergent angiosperm taxa (Hiscock and Allen, 2008).

Production of this secretion is already seen before flower opening and reaches a maximum level 1 d after anthesis, concomitant with pollen tube passage. However, the production of secretion is independent of pollination as it occurs in the same way and at the same time in pollinated and unpollinated flowers. This contrasts with higher angiosperms, in which pollen tube growth in the style triggers starch degradation (Herrero and Dickinson, 1979; Gonzalez et al., 1996). Interestingly, however, this production of secretion is very similar to the behaviour of the obturator in which secretion occurs at a particular time of development independent of pollination (Herrero and Arbeloa, 1989; Arbeloa and Herrero, 1991). In A. cherimola a primitive obturator, formed by the protuberance of the placenta, continuous with and with the same cytohistological features as the secretory papillae, appears to be present in the ovary. Similar structures have been described in other ancient angiosperm lineages, such as in species of the Magnoliaceae that show a funicular outgrowth with papillose cells (Matsui et al., 1993; Umeda et al., 1994), in Schisandraceae (Lyew et al., 2007), Lauraceae (Sedgley and Annells, 1981) and in the monocot Ornithogalum caudatum (Tilton and Horner, 1980). In A. cherimola the fact that secretion is present right from anthesis at pollination provides an adequate substrate for rapid pollen tube growth. This contrasts with longer times for pollen tube growth reported in other species, which are related to waiting times in order to reach the phase where secretion is produced in the pistilar structures (Herrero and Arbeloa, 1989; Herrero, 2000, 2003).

Following the production of secretion, callose is layered in the papillar secretory structures in the same way reported in the obturator of peach (Arbeloa and Herrero, 1987), perhaps protecting this area and fulfilling what has been considered as a prophylactic role (Heslop-Harrison, 1999, 2000).

Pistil constraint to pollen tube growth

Pollen tube growth proceeds rapidly and, within 1 d of pollination, the pollen tubes reach the hood-shaped ovule, which has been considered as a plesiomorphic trait in angiosperms (Soltis et al., 2005). Relatively rapid pollen tube growth has also been found in other members of ancient angiosperm clades (Bernhardt et al., 2003; Sage and Sampson, 2003; Koehl et al., 2004; Hristova et al., 2005; Williams, 2008, 2009) and contrasts with the slow growth of pollen tubes in gymnosperms (Gelbart and Von Aderkas, 2002). Both the pollen tube growth rate of A. cherimola (480 µm h−1) and the length of the pollen tube pathway (1·83) mm are in the range described for basal grade angiosperms (Williams, 2008: approx. 80–600 µm h−1 and <0·5 to approx. 15 mm long). Although in some derived angiosperms delayed fertilization has also been recorded (Sogo and Tobe, 2005, 2006a, b), an evolutionary trend towards rapid pollen tube growth in seed plant pollen has been proposed (Pettitt, 1982; Williams, 2008) where the development of callose plugs in pollen tubes could have played a major role (Williams, 2008). Differences in timing also appear to be related to differences in maturation of the pistil (Herrero and Arbeloa, 1989; Sogo and Tobe, 2005, 2006a, b) and to a requirement for male–female synchrony (Herrero, 2003).

Several pollen grains germinate on the stigma, but only one pollen tube reaches the ovule and achieves fertilization. Although pollen grains germinate freely at the stigma and direct their growth towards the semi-open suture line, a clear restriction and reduction in the number of pollen tubes occur at this point of entrance in the short stylar canal. Only 1–3 pollen tubes penetrate the style. This reduction in the number of pollen tubes may be related to the limited space available with only a narrow papillar secretory area that paves, along the semi-open suture line, the carpel margins. Pollen competition and selection appears to be a common feature shared by most angiosperms (Mulcahy, 1979; Hormaza and Herrero, 1992, 1996) and it is usually reflected by a reduction in the number of pollen tubes that continue to grow in the style (Sedgley, 1977; Cruzan, 1990; Hormaza and Herrero, 1996). However, results herein show that in A. cherimola the main restriction point appears at the stigma–style interface. This behaviour should be investigated in other early-divergent angiosperms, but interesting recent work in Amborella trichopoda (Williams, 2009) shows a very similar behaviour. A. cherimola has a semi-open stylar canal similar to that described in Amborella trichopoda (Williams, 2009) and in A. cherimola only the carpel margins are layered with secretory papillae, paving a narrow way for the few pollen tubes growing in the style. A semi-open stylar canal is a common feature found in other ancient angiosperms (Endress and Igersheim, 2000) and it would be interesting to evaluate if papillar secretory cells restricted to the margins also provide a similar pollen restriction mechanism, in contrast to the typical pollen tube attrition recorded in the style in evolutionarily derived angiosperms. If this is so, during angiosperm evolution, the arena for pollen competition would have changed from the stigma to within the style and, consequently, pollen competition in the style could be considered an innovation in evolutionarily derived clades of flowering plants.

Post-fertilization events

Three days after pollination callose is layered in the nucellus under the embryo sac micropylar pole only in the ovules of pollinated flowers that appear to have been fertilized. By contrast, in ovules of unpollinated flowers, deposition of callose in vascular bundles was observed 6 d after pollination, suggesting impending ovule abortion. This has been shown in other species and explained in terms of blockage of metabolite translocation (Pimienta and Polito, 1982; Herrero and Arbeloa, 1989; Rodrigo and Herrero, 1998).

The presence of starch grains has been reported in mature embryo sacs in some ancient angiosperm lineages (Cook, 1902; Kimoto and Tobe, 2001; Friedman, 2008), including species in the Annonaceae (reviewed in Svoma, 1998b), and also in higher angiosperms (Evans, 1919; Maheshwari, 1950). However, the abundance and large size of the starch grains observed in the present study are striking. Recent results in Hydatellaceae (Friedman, 2008; Rudall et al., 2008), a family recently recognized among early-divergent extant angiosperms (Saarela et al., 2007), show a maternal seed-provisioning strategy similar to that observed in gymnosperms. The provision of starch grains reported here in the embryo sac before fertilization in A. cherimola could respond to a similar plesiomorphic strategy. Through a different accumulation pattern, the accumulation of starch reserves in either the sporophytic or the gametophytic tissue would constitute an accumulation of reserves before fertilization to support early post-fertilization processes.

Although zygote cell division does not start until 8 d after pollination, endosperm cell division starts 3 d after pollination. Division of the endosperm is bipolar, giving rise to a large cell close to the zygote and a small cell full of starch at the chalazal end. This situation persists during the first endosperm divisions. Although this behaviour in relation to starch accumulation has not been reported previously, a bipolar endosperm cellular division has been shown in several species of the Winteraceae (a sister group to Annonaceae in the Magnoliales) such as Drimys winteri (Bhandari and Venkatar, 1968; Floyd and Friedman, 2000), Pseudowintera axillaries (Sampson, 1963, cited in Bhandari and Venkatar, 1968) and Zygogynum bailloni (Swamy, 1952). The presence of a similar cellular endosperm with unequal division has also been reported in other ancient angiosperms (Floyd and Friedman, 2000; Tobe et al., 2000) and seems to be a plesiomorphic feature in angiosperms. The prominence of endosperm development in A. cherimola contrasts with an underdeveloped embryo that in the mature seed is embedded in abundant ruminate endosperm, similar to other species of the Annonaceae (Corner, 1949; Svoma, 1998b), where this slight embryo development has been postulated as an ancestral feature (Hayat, 1963; Finch-Savage and Leubner-Metzger, 2006).

CONCLUDING REMARKS

Flowers of A. cherimola have a number of ancestral characteristics of angiosperms such as the semi-open simple carpel, hood-shaped ovule, cellular endosperm and seed type with a rudimentary embryo. However, this primitive carpel hosts a support–constrain strategy for pollen tube growth conserved in phylogenetically derived angiosperm lineages. Support is provided by the continuous secretory papillar carpet that paves the pollen tube pathway and that provides evidence for a common ontogenetic origin for this tissue as well as for a conserved basipetal maturation that encompasses pollen tube growth. Constraint and restriction in the number of pollen tubes occurs at the stigma–style interface and is mediated by a dramatic reduction in the secretory surface available for pollen tube growth from the stigma to the margins of the semi-open stylar canal. It will be of interest to evaluate in other ancient lineages of angiosperms with a similar pistil anatomy if this extramural pollen competition is conserved as compared with the stylar intramural competition in modern angiosperms.

ACKNOWLEDGEMENTS

This work was supported by the Spanish Ministry of Education (Project Grants AGL2004-02290/AGR, AGL2006-13529 and AGL2007-60130/AGR), GIC-Aragón 43, Junta de Andalucía (AGR2742) and the European Union under the INCO-DEV programme (Contract 015100). J.L. was supported by a grant from Junta de Andalucía.

LITERATURE CITED

  1. APG II. An update of the Angiosperm Phylogeny Group classification for the orders and families of flowering plants: APG II. Botanical Journal of the Linnean Society. 2003;141:399–436. [Google Scholar]
  2. Arbeloa A, Herrero M. The significance of the obturator in the control of pollen tube entry into the ovary in peach (Prunus persica) Annals of Botany. 1987;60:681–685. [Google Scholar]
  3. Arbeloa A, Herrero M. Development of the ovular structures in peach [Prunus-persica (L.) Batsch] New Phytologist. 1991;118:527–533. [Google Scholar]
  4. Bernhardt P, Sage T, Weston P, et al. The pollination of Trimenia moorei (Trimeniaceae): floral volatiles, insect/wind pollen vectors and stigmatic self-incompatibility in a basal angiosperm. Annals of Botany. 2003;92:445–458. doi: 10.1093/aob/mcg157. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bhandari NN, Venkatar R. Embryology of Drimys winteri. Journal of the Arnold Arboretum. 1968;49:509–525. [Google Scholar]
  6. Carvalho R, Weber AC. Biologia floral de Unonopsis guatterioides (A. D.C.) R.E. Fr., uma Annonaceae polinizada por Euglossini. Revista Brasileira de Botânica. 2000;23:421–425. [Google Scholar]
  7. Chatrou LW, Rainer H, Mass PJM. Annonaceae. In: Smith N, Mori SA, Henderson A, Steveson DW, Heald SV, editors. Flowering plants of the neotropics. Princeton, NJ: Princeton University Press; 2004. pp. 18–20. [Google Scholar]
  8. Cook MT. Development of the embryo-sac and embryo of Castalia odorata and Nymphaea advena. Bulletin of the Torrey Botanical Club. 1902;29:211–220. [Google Scholar]
  9. Corner EJH. The Annonaceous seed and its four integuments. New Phytologist. 1949;48:332–364. [Google Scholar]
  10. Cruzan MB. Pollen–pollen and pollen–style interactions during pollen-tube growth in Erythronium grandiflorum (Liliaceae) American Journal of Botany. 1990;77:116–122. [Google Scholar]
  11. Currier HB. Callose substance in plant cells. American Journal of Botany. 1957;44:478–488. [Google Scholar]
  12. Decraene LPR, Smets E. The floral development of Popowia whitei (Annonaceae) Nordic Journal of Botany. 1990;10:411–420. [Google Scholar]
  13. Dudley ME, Jacobs TW, Long SR. Microscopic studies of cell divisions induced in alfalfa roots by Rhizobium meliloti. Planta. 1987;171:289–301. doi: 10.1007/BF00398674. [DOI] [PubMed] [Google Scholar]
  14. Endress PK, Igersheim A. Gynoecium structure and evolution in basal angiosperms. International Journal of Plant Sciences. 2000;161:S211–S223. doi: 10.1086/314241. [DOI] [PubMed] [Google Scholar]
  15. Escobar-Restrepo JM, Huck N, Kessler S, et al. The FERONIA receptor-like kinase mediates male–female interactions during pollen tube reception. Science. 2007;317:656–660. doi: 10.1126/science.1143562. [DOI] [PubMed] [Google Scholar]
  16. Evans A. Embryo sac and embryo of Pentstemon secundiflorus. Botanical Gazette. 1919;67:427–437. [Google Scholar]
  17. Feder N, O'Brien TP. Plant microtechnique: some principles and new methods. American Journal of Botany. 1968;55:123–142. [Google Scholar]
  18. Finch-Savage WE, Leubner-Metzger G. Seed dormancy and the control of germination. New Phytologist. 2006;171:501–523. doi: 10.1111/j.1469-8137.2006.01787.x. [DOI] [PubMed] [Google Scholar]
  19. Floyd SK, Friedman WE. Evolution of endosperm developmental patterns among basal flowering plants. International Journal of Plant Sciences. 2000;161:S57–S81. [Google Scholar]
  20. Friedman WE. Embryological evidence for developmental lability during early angiosperm evolution. Nature. 2006;441:337–340. doi: 10.1038/nature04690. [DOI] [PubMed] [Google Scholar]
  21. Friedman WE. Hydatellaceae are water lilies with gymnospermous tendencies. Nature. 2008;453:94–97. doi: 10.1038/nature06733. [DOI] [PubMed] [Google Scholar]
  22. Friedman WE, Ryerson KC. Reconstructing the ancestral female gametophyte of angiosperms: insights from Amborella and other ancient lineages of flowering plants. American Journal of Botany. 2009;96:129–143. doi: 10.3732/ajb.0800311. [DOI] [PubMed] [Google Scholar]
  23. Friedman WE, Williams JH. Developmental evolution of the sexual process in ancient flowering plant lineages. Plant Cell. 2004;16:S119–S132. doi: 10.1105/tpc.017277. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Friedman WE, Madrid EN, Williams JH. Origin of the fittest and survival of the fittest: relating female gametophyte development to endosperm genetics. International Journal of Plant Sciences. 2008;169:79–92. [Google Scholar]
  25. Gelbart G, Von Aderkas P. Ovular secretions as part of pollination mechanisms in conifers. Annals of Forest Science. 2002;59:345–357. [Google Scholar]
  26. Gerlach D. A rapid safranin-crystal violet-light green staining sequence for paraffin sections of plant materials. Stain Technology. 1969;44:210–211. [PubMed] [Google Scholar]
  27. Gonzalez MV, Coque M, Herrero M. Pollen–pistil interaction in kiwifruit (Actinidia deliciosa; Actinidiaceae) American Journal of Botany. 1996;83:148–154. [Google Scholar]
  28. de Graaf BHJ, Derksen JWM, Mariani C. Pollen and pistil in the progamic phase. Sexual Plant Reproduction. 2001;14:41–55. [Google Scholar]
  29. Hayat AM. Morphology of seed germination and seedling in Annona squamosa. Botanical Gazette. 1963;124:360–362. [Google Scholar]
  30. Herrero M. Changes in the ovary related to pollen tube guidance. Annals of Botany. 2000;85:79–85. [Google Scholar]
  31. Herrero M. Male and female synchrony and the regulation of mating in flowering plants. Philosophical Transactions of the Royal Society of London Series B-Biological Sciences. 2003;358:1019–1024. doi: 10.1098/rstb.2003.1285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Herrero M, Arbeloa A. Influence of the pistil on pollen-tube kinetics in peach (Prunus persica) American Journal of Botany. 1989;76:1441–1447. [Google Scholar]
  33. Herrero M, Dickinson HG. Pollen–pistil incompatibility in Petunia hybrida: changes in the pistil following compatible and incompatible intraspecific crosses. Journal of Cell Science. 1979;36:1–18. doi: 10.1242/jcs.36.1.1. [DOI] [PubMed] [Google Scholar]
  34. Herrero M, Hormaza JI. Pistil strategies controlling pollen tube growth. Sexual Plant Reproduction. 1996;9:343–347. [Google Scholar]
  35. Heslop-Harrison J. The structure and prophylactic role of the angiosperm embryo sac and its associated tissues. Zea mays as a model. Protoplasma. 1999;209:256–272. [Google Scholar]
  36. Heslop-Harrison Y. Pollen–stigma interaction – pollen-tube penetration in Crocus. Annals of Botany. 1977;41:913–922. [Google Scholar]
  37. Heslop-Harrison Y. Control gates and micro-ecology: the pollen–stigma interaction in perspective. Annals of Botany. 2000;85:5–13. [Google Scholar]
  38. Heslop-Harrison Y, Shivanna KR. Receptive surface of angiosperm stigma. Annals of Botany. 1977;41:1233–1258. [Google Scholar]
  39. Higashiyama T, Hamamura Y. Gametophytic pollen tube guidance. Sexual Plant Reproduction. 2008;21:17–26. [Google Scholar]
  40. Hiscock SJ, Allen AM. Diverse cell signalling pathways regulate pollen-stigma interactions: the search for consensus. New Phytologist. 2008;179:286–317. doi: 10.1111/j.1469-8137.2008.02457.x. [DOI] [PubMed] [Google Scholar]
  41. Hormaza JI, Herrero M. Pollen selection. Theoretical and Applied Genetics. 1992;83:663–672. doi: 10.1007/BF00226682. [DOI] [PubMed] [Google Scholar]
  42. Hormaza JI, Herrero M. Dynamics of pollen tube growth under different competition regimes. Sexual Plant Reproduction. 1996;9:153–160. [Google Scholar]
  43. Hristova K, Lam M, Field T, Sage TL. Transmitting tissue ECM distribution and composition, and pollen germinability in Sarcandra glabra and Chloranthus japonicus (Chloranthaceae) Annals of Botany. 2005;96:779–791. doi: 10.1093/aob/mci235. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Hughes J, McCully ME. Stain Technology 50. 1975 doi: 10.3109/10520297509117082. The use of an optical brightener in the study of plant structure 319. [DOI] [PubMed] [Google Scholar]
  45. Johansen DA. Plant microtechnique. New York: McGraw-Hill; 1940. [Google Scholar]
  46. Kiill LHP, Costa JG. Biologia floral e sistema de reprodução de Annona squamosa L. (Annonaceae) na região de Petrolina-PE. Ciência Rural. 2003;533:851–856. [Google Scholar]
  47. Kimoto Y, Tobe H. Embryology of Laurales: a review and perspectives. Journal of Plant Research. 2001;114:247–267. [Google Scholar]
  48. Koehl V, Thien LB, Heij EG, Sage TL. The causes of self-sterility in natural populations of the relictual angiosperm. Illicium floridanum (Illiciaceae). Annals of Botany. 2004;94:43–50. doi: 10.1093/aob/mch113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Leins P, Erbar C. Early floral developmental studies in Annonaceae. In: Morawetz W, Winkler H, editors. Reproductive morphology in Annonaceae. Vienna: Österreich Akademie der Wissenschaften; 1996. pp. 1–27. [Google Scholar]
  50. Linskens HF. Incompatibility in Petunia. Proceedings of the Royal Society of London Series B-Biological Sciences. 1975;188:299–311. [Google Scholar]
  51. Linskens HF, Esser K. Über eine spezifische Anfärbung der Pollen-Schläuche und die Zahl Kallosepfropen nach Selbstung und Fremdung. Naturwissenschaften. 1957;44::16. [Google Scholar]
  52. Lora J, Herrero M, Hormaza JI. The coexistence of bicellular and tricellular pollen in Annona cherimola (Annonaceae): implications for pollen evolution. American Journal of Botany. 2009a;96:802–808. doi: 10.3732/ajb.0800167. [DOI] [PubMed] [Google Scholar]
  53. Lora J, Testillano PS, Risueño MC, Hormaza JI, Herrero M. BMC Plant Biology. 2009b;9:129–139. doi: 10.1186/1471-2229-9-129. Pollen development in Annona cherimola Mill. (Annonaceae). Implications for the evolution of aggregated pollen. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Lyew J, Li Z, Liang-Chen Y, Yi-Bo L, Sage TL. Pollen tube growth in association with a dry-type stigmatic transmitting tissue and extragynoecial compitum in the basal angiosperm Kadsura longipedunculata (Schisandraceae) American Journal of Botany. 2007;94:1170–1182. doi: 10.3732/ajb.94.7.1170. [DOI] [PubMed] [Google Scholar]
  55. Maheshwari P. An introduction to the embryology of angiosperms. New York: McGraw-Hill; 1950. [Google Scholar]
  56. Matsui M, Imaichi R, Kato M. Ovular development and morphology in some Magnoliaceae species. Journal of Plant Research. 1993;106:297–304. [Google Scholar]
  57. Mulcahy DL. The rise of the angiosperms: a genecological factor. Science. 1979;206:20–23. doi: 10.1126/science.206.4414.20. [DOI] [PubMed] [Google Scholar]
  58. Nicholas JR, Gates PJ, Grierson D. The use of fluorescence microscopy to monitor root development in micropropagated explants. Journal of Horticultural Science. 1986;61:417–421. [Google Scholar]
  59. Norman EM. Reproductive biology of Deeringothamnus rugelii and D. pulchellus (Annonaceae) Taxon. 2003;52:547–555. [Google Scholar]
  60. Norman EM, Clayton DE, Rice K, Cochran S. Reproductive biology of two Florida pawpaws: Asimina obovata and Asimina pygmaea (Annonaceae) Bulletin of the Torrey Botanical Club. 1986;113:16–22. [Google Scholar]
  61. Norman EM, Rice K, Cochran S. Reproductive biology of Asimina parviflora (Annonaceae) Bulletin of the Torrey Botanical Club. 1992;119:1–5. [Google Scholar]
  62. Orban I, Bouharmont J. Reproductive biology of Nymphaea capensis Thunb. var. zanzibariensis (Casp) Verdc (Nymphaeaceae) Botanical Journal of the Linnean Society. 1995;119:35–43. [Google Scholar]
  63. Pettitt JM. Ultrastructural and immuno-cytochemical demonstration of gametophytic proteins in the pollen-tube wall of the primitive gymnosperm Cycas. Journal of Cell Science. 1982;57:189–213. doi: 10.1242/jcs.57.1.189. [DOI] [PubMed] [Google Scholar]
  64. Pimienta E, Polito VS. Ovule abortion in nonpareil almond (Prunus dulcis [Mill.] D.A. Webb) American Journal of Botany. 1982;69:913–920. [Google Scholar]
  65. Pontieri V, Sage TL. Evidence for stigmatic self-incompatibility, pollination induced ovule enlargement and transmitting tissue exudates in the paleoherb, Saururus cernuus L. (Saururaceae) Annals of Botany. 1999;84:507–519. [Google Scholar]
  66. Popenoe H. Lost crops of the Incas: little known plants of the Andes with promise of worldwide cultivation. Washington, DC: National Academy Press; 1989. [Google Scholar]
  67. Rea AC, Nasrallah JB. Self-incompatibility systems: barriers to self-fertilization in flowering plants. International Journal of Developmental Biology. 2008;52:627–636. doi: 10.1387/ijdb.072537ar. [DOI] [PubMed] [Google Scholar]
  68. Rodrigo J, Herrero M. Influence of intraovular reserves on ovule fate in apricot (Prunus armeniaca L.) Sexual Plant Reproduction. 1998;11:86–93. [Google Scholar]
  69. Rudall PJ, Bateman RM. Developmental bases for key innovations in the seed-plant microgametophyte. Trends in Plant Science. 2007;12:317–326. doi: 10.1016/j.tplants.2007.06.004. [DOI] [PubMed] [Google Scholar]
  70. Rudall PJ, Remizowa MV, Beer AS, et al. Comparative ovule and megagametophyte development in Hydatellaceae and water lilies reveal a mosaic of features among the earliest angiosperms. Annals of Botany. 2008;101:941–956. doi: 10.1093/aob/mcn032. [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Rudall PJ, Remizowa MV, Prenner G, Prychid CJ, Tuckett RE, Sokoloff DD. Nonflowers near the base of extant angiosperms? Spatiotemporal arrangement of organs in reproductive units of Hydatellaceae and its bearing on the origin of the flower. American Journal of Botany. 2009;96:67–82. doi: 10.3732/ajb.0800027. [DOI] [PubMed] [Google Scholar]
  72. Saarela JM, Rai HS, Doyle JA, et al. Hydatellaceae identified as a new branch near the base of the angiosperm phylogenetic tree. Nature. 2007;446:312–315. doi: 10.1038/nature05612. [DOI] [PubMed] [Google Scholar]
  73. Sabatini DD, Bensch K, Barrnett RJ. Cytochemistry and electron microscopy – preservation of cellular ultrastructure and enzymatic activity by aldehyde fixation. Journal of Cell Biology. 1963;17:19–58. doi: 10.1083/jcb.17.1.19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Sage TL, Sampson FB. Evidence for ovarian self-incompatibility as a cause of self-sterility in the relictual woody angiosperm, Pseudowintera axillaris (Winteraceae) Annals of Botany. 2003;91:807–816. doi: 10.1093/aob/mcg085. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Sage TL, Hristova-Sarkovski K, Koehl V, et al. Transmitting tissue architecture in basal-relictual angiosperms: implications for transmitting tissue origins. American Journal of Botany. 2009;96:183–206. doi: 10.3732/ajb.0800254. [DOI] [PubMed] [Google Scholar]
  76. Sampson FB. The floral morphology of Pseudowintera, the New Zealand member of the vesselless Winteraceae. Phytomorphology. 1963;13:403–423. [Google Scholar]
  77. Schroeder CA. Pollination of cherimoya. California Avocado Society Yearbook. 1971;44:119–122. [Google Scholar]
  78. Sedgley M. Reduced pollen tube growth and the presence of callose in the pistil of the male floral stage of the avocado. Scientia Horticulturae. 1977;7:27–36. [Google Scholar]
  79. Sedgley M, Annells CM. Flowering and fruit-set response to temperature in the avocado cultivar Hass. Scientia Horticulturae. 1981;14:27–33. [Google Scholar]
  80. Sogo A, Tobe H. Intermittent pollen-tube growth in pistils of alders (Alnus) Proceedings of the National Academy of Sciences USA. 2005;102:8770–8775. doi: 10.1073/pnas.0503081102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Sogo A, Tobe H. Mode of pollen-tube growth in pistils of Myrica rubra (Myricaceae): a comparison with related families. Annals of Botany. 2006a;97:71–77. doi: 10.1093/aob/mcj015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  82. Sogo A, Tobe H. Delayed fertilization and pollen-tube growth in pistils of Fagus japonica (Fagaceae) American Journal of Botany. 2006b;93:1748–1756. doi: 10.3732/ajb.93.12.1748. [DOI] [PubMed] [Google Scholar]
  83. Soltis DE, Soltis PS, Endress PK, Chase MW. Phylogeny and evolution of angiosperms. Sunderland, MA: Sinauer Associates Incorporated; 2005. [Google Scholar]
  84. Svoma E. Studies on the embryology and gynoecium structures in Drimys winteri (Winteraceae) and some Annonaceae. Plant Systematics and Evolution. 1998a;209:205–229. [Google Scholar]
  85. Svoma E. Seed morphology and anatomy in some Annonaceae. Plant Systematics and Evolution. 1998b;209:177–204. [Google Scholar]
  86. Swamy BGL. Some aspects in the embryology of Zygogynum bailloni V. Tiegh. Proceedings of the National Institute of Sciences of India. 1952;18:399–406. [Google Scholar]
  87. Thien LB, Sage TL, Jaffre T, et al. The population structure and floral biology of Amborella trichopoda (Amborellaceae) Annals of the Missouri Botanical Garden. 2003;90:466–490. [Google Scholar]
  88. Thien LB, Bernhardt P, Devall MS, et al. Pollination biology of basal angiosperms (ANITA grade) American Journal of Botany. 2009;96:166–182. doi: 10.3732/ajb.0800016. [DOI] [PubMed] [Google Scholar]
  89. Tilton VR, Horner HT. Stigma, style, and obturator of Ornithogalum caudatum (Liliaceae) and their function in the reproductive process. American Journal of Botany. 1980;67:1113–1131. [Google Scholar]
  90. Tobe H, Jaffré T, Raven PH. Embryology of Amborella (Amborellaceae): descriptions and polarity of character states. Journal of Plant Research. 2000;113:271–280. [Google Scholar]
  91. Umeda A, Imaichi R, Kato M. Ovular development and morphology of the outer integument of Magnolia grandiflora (Magnoliaceae) American Journal of Botany. 1994;81:361–367. [Google Scholar]
  92. Vithanage HIMV. Pollen stigma interactions – development and cyto-chemistry of stigma papillae and their secretions in Annona squamosa L. (Annonaceae) Annals of Botany. 1984;54:153–167. [Google Scholar]
  93. Williams JH. Novelties of the flowering plant pollen tube underlie diversification of a key life history stage. Proceedings of the National Academy of Sciences USA. 2008;105:11259–11263. doi: 10.1073/pnas.0800036105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  94. Williams JH. Amborella trichopoda (Amborellaceae) and the evolutionary developmental origins of the angiosperm progamic phase. American Journal of Botany. 2009;96:144–165. doi: 10.3732/ajb.0800070. [DOI] [PubMed] [Google Scholar]
  95. Williams JH, Friedman WE, Arnold ML. Developmental selection within the angiosperm style: using gamete DNA to visualize interspecific pollen competition. Proceedings of the National Academy of Sciences USA. 1999;96:9201–9206. doi: 10.1073/pnas.96.16.9201. [DOI] [PMC free article] [PubMed] [Google Scholar]

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