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(American Journal of Botany. 2009;96:183-206.) doi: 10.3732/ajb.0800254 © 2009 Botanical Society of America, Inc. |
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Special Invited Papers |
2 Department of Ecology and Evolutionary Biology, University of Toronto, Toronto, Ontario, Canada M5S 3B2 3 Department of Biology, Saint Louis University, St. Louis, Missouri, USA 63103-2010 4 Royal Botanic Gardens, Mrs. Macquaries Road, Sydney, Australia 2000, Australia
Received for publication 23 July 2008. Accepted for publication 24 November 2008.
ABSTRACT
Carpel transmitting tissue is a major floral innovation that is essential for angiosperm success. It facilitates the rapid adhesion, hydration, and growth of the male gametophyte to the female gametophyte. As well, it functions as a molecular screen to promote male gametophytic competition and species-specific recognition and compatibility. Here, we characterize the transmitting tissue extracellular matrix (ECM) and pollen tube growth in basal-relictual angiosperms and test the hypothesis that a freely flowing ECM (wet stigma) was ancestral to a cuticle-bound ECM (dry stigma). We demonstrate that the most recent common ancestor of extant angiosperms produced an ECM that was structurally and functionally equivalent to a dry stigma. Dry stigmas are composed of a cuticle and primary wall that contains compounds that facilitate the adhesion and growth of the male gametophyte. These compounds include methyl-esterified homogalacturonans, arabinogalactan-proteins, and lipids. We propose that transmitting tissue evolved in concert with an increase in cuticle permeability that resulted from modifications in the biosynthesis and secretion of fatty acids needed for cuticle construction. Increased cuticle permeability exposed the male gametophyte to pre-existing molecules that enabled rapid male gametophyte adhesion, hydration, and growth as well as species-specific recognition and compatibility.
Key Words: arabinogalactan-proteins • cuticle • dry stigma • homogalacturonans • pollen-carpel interactions • pollination droplet • transmitting tissue • wet stigma
The evolutionary success and consequent persistence of the ovule-seed has been closely tied to modifications of tissues that optimized capture, retention, germination, and transmission of the male gametophyte to the site of fertilization. Features of ovules that have served to increase the probability of passive pollen capture and retention include the funnel-shaped projection from the nucellus, the salpinx or lagenostome in lyginopterid pteridosperms (Camp and Hubbard, 1963
), the tubular-shaped integumentary micropyle, and the secretion of the pollination droplet (Dilcher, 1979; Taylor and Millay, 1979; Dilcher, 1979; Crane, 1985; Buchmann et al., 1989; Owens, et al., 1998; Gelbart and von Aderkas, 2002; Stockey and Rothwell, 2003). Active mechanisms of pollen capture and retention in gymnosperms have included sealing of the micropyle or pollen chamber by a plug or proliferation of tissues surrounding the megagametophyte (Owens et al., 1998; Rothwell and Stockey, 2002; Stockey and Rothwell, 2003).
The evolution of the carpel and carpel closure resulted in the elimination of the ovule as the site of pollen capture and initial pollen tube growth to the female gametophyte. Therefore, the evolution of carpel transmitting tissue that operates in supporting growth of the male gametophyte to the female gametophyte was crucial for angiosperm success. Given the pivotal role of transmitting tissue for completion of the fertilization process as well as promoting rapid reproduction in angiosperms (Stebbins, 1976), male gametophytic competition and selection (Mulcahy, 1975; Willson and Burley, 1983), and the union of appropriately matched gametes through mechanisms that confer species recognition and compatibility (de Nettancourt, 1997, 2001), the evolutionary processes giving rise to transmitting tissue and its ancestral features have been long-standing topics of interest. Consequently, numerous ideas have been formulated regarding the histogenetic origins of transmitting tissue and traits of the associated extracellular matrix (ECM). The most common hypothesis predicts that the site of production of a gymnosperm-like pollination droplet shifted from the ancestral gymnosperm ovule to early carpellary structures through the development of glandular hairs on the margins and ventral epidermis of the carpel progenitor (Thomas, 1931, 1934; Bailey and Swamy, 1951; Lloyd and Wells, 1992; Frohlich and Parker, 2000; Frame, 2003a, b; Frohlich and Chase, 2007). The observation that nonovular organs such as ovuliferous scales of some gymnosperms are involved in pollen capture (Haines et al., 1984; Owens et al., 1998) is often used as support for this spatial shift (Lloyd and Wells, 1992; Frohlich and Parker, 2000; Frame, 2003a, b; Frohlich and Chase, 2007). An alternative hypothesis states that transmission tissue was derived from homologues of the bennettitalean interseminal scales that are sterilized ovules (Cornet, 1989). Such a scenario has been viewed as intriguing because it prevents the need for the transfer of function to nonovular organs with no "prior experience" (Cornet, 1989). However, transfer of function to nonovular organs with no prior experience may not be a significant developmental obstacle given the role of the ovuliferous scale in pollen capture in gymnosperms and ectopic germination of pollen on leaves and stems of plants deficient in cuticle synthesis (Lolle and Cheung, 1993; Sieber et al., 2000). Alternatively, mutagenic sterilization of ovules can give rise to carpelloid structures with functional transmitting tissue to include marginal papillate and ventral epidermal cells (Angenent et al., 1995).
The envisioned ancestral condition of the ventral and marginal glandular epidermis with its associated pollination droplet has been compared to the wet stigma found in extant angiosperms (Lloyd and Wells, 1992; Frame, 2003a, b). The extracellular matrix (ECM) of both wet and dry stigmas is contained by a cuticle and associated protein layer with esterase activity (pellicle; Mattsson et al., 1974; Heslop-Harrison and Heslop-Harrison, 1975; Mattsson et al., 1974; Heslop-Harrison and Shivanna, 1977). However, in a dry stigma, the esterase-positive pellicle and cuticle remain intact in the functional phase, whereas they become ruptured in the wet stigma thereby resulting in a freely flowing secretion (Heslop-Harrison and Shivanna, 1977; Shivanna and Sastri, 1981). The pellicle-cuticle of the dry stigma provides a well-defined focal point of interaction between a pollen grain and stigmatic ECM molecules. This trait, viewed to be strictly correlated with a derived character state of tricellular pollen (Hiscock and Allen, 2008), is believed to provide more site-specific control over species-specific compatible pollen grain adhesion, hydration, and germination than that offered by a wet stigma or pollination droplet of gymnosperms (Dickinson, 1995; Edlund et al., 2004; Swanson et al., 2004; Hiscock and Allen, 2008). This enhanced selectivity over male gametophyte growth conferred by the dry stigma appears to be true in most, but not all species (Heslop-Harrison, 1982). And mechanisms have evolved in gymnosperms with pollination droplets that prevent the growth of unrelated pollen (Runions et al., 1999). The evolution of dry stigmas from wet stigmas has been proposed to be a significant step in the evolution of angiosperms (Dickinson, 1995).
The timing of carpel closure with respect to the origins of the ancestral stigma has been an open question, and the glandular ventral and marginal stigma has been suggested to have appeared either before or concurrently with carpel closure (Whitehouse, 1950; Lloyd and Wells, 1992). In either case, the ancestral stigma and its associated "pollination droplet," or freely flowing ECM, have been proposed to have functioned not only in pollen capture and pollen tube growth, but also as a protonectary to attract pollinators prior to, and after carpel closure before the evolution of nectaries (Bernhardt and Thien, 1987; Gottsberger, 1988; Lloyd and Wells, 1992; Endress and Igersheim, 2000a; Frohlich and Parker, 2000; Frame, 2003a, b; Frohlich and Chase, 2007; Labandeira et al., 2007). After carpel closure, the ventral glandular epidermis and its associated freely flowing ECM at the site of carpel closure, which would have no longer been available for pollen capture, have been presumed to have not only continued to contribute to pollen tube transmission to the ovules but to carpel closure as well (Lloyd and Wells, 1992; Endress and Igersheim, 2000a; Frame, 2003a, b). And, while both the site of carpel closure and the marginal stigma have been viewed as the ancestral location of male gametophytic competition and selection and recognition of self through mechanisms of self-incompatibility (SI; Whitehouse, 1950; Zavada, 1984; Zavada and Taylor, 1986; Bernhardt and Thien, 1987; Lloyd and Wells, 1992; Endress and Igersheim, 2000a; de Nettancourt, 2001), these processes have also been proposed to have functioned prior to carpel closure either within the ovule (Kenrick et al., 1986) or on early carpellary structures in association with the evolution of penetrative siphonogamy (Bell, 1995).
The recently resolved patterns of extant basal-relictual angiosperm relationships (summarized by Stevens, 2001) provide new opportunities for phylogenetic reconstruction of the reproductive characters potentially present in early angiosperms using comparative work on extant lineages that branch at or near the base of the angiosperm tree (Friis et al., 2000; Endress, 2001, 2004). The ability to use these taxa to test hypotheses about the early evolution of the transmitting tissue and associated ECM and its role in pollen tube growth, carpel closure, and insect attraction requires, as a first step, an understanding of the spatial distribution of transmitting tissue, cellular features of transmitting tissue and associated ECM, and pollen–carpel interactions therein. In contrast to the hypothesis that a freely flowing stigmatic ECM represents the ancestral condition in angiosperms, studies on species within the magnoliid family Saururaceae (Pontieri and Sage, 1999; Pontieri, 2004), three species from the ANITA paraphyletic grade, Amborella trichopoda Baill. (Amborellaceae; Thien et al., 2003), Trimenia moorei (Oliv.) Philipson (Trimeniaceae; Bernhardt et al., 2003), Kadsura longipedunculata Finet and Gagnep (Schisandraceae; Lyew et al., 2007), and two species, Chloranthus japonicus Siebold and Sarcandra glabra (Thunb.) Nakai, from the family Chloranthaceae (Hristova et al., 2005), which has been variously placed closer to the monocots, eudicots, and magnoliids than to any member of the "ANITA" grade (Zanis et al., 2002; Eklund et al., 2004; Moore et al., 2007), have demonstrated that stigmas are of the dry-type as indicated by the presence of a pellicle overlying the intact cuticle at stigma receptivity. As well, all these species have bicellular pollen, thus calling into question broad generalizations linking pollen morphology with stigmatic ECM type (Hiscock and Allen, 2008). Studies examining the pollination biology of Amborella (Thien et al., 2003), Trimenia (Bernhardt et al., 2003), and Saururus cernuus L. (Thien et al., 1994) have documented that the dry-type stigma does not function as a nectary to attract pollinators. In these species, pollen is the primary attractant. When the distribution of transmitting tissue beyond the stigma is considered, data show that transmitting tissue is not confined to the ventral epidermis at the site of carpel closure in Trimenia, Chloranthus, and Sarcandra as predicted. Rather, in these species, pollen tubes display penetrative siphonogamy en route to the ovary locule following stigmatic germination by growing within the stigmatic ECM (Bernhardt et al., 2003; Hristova et al., 2005). Similar patterns of pollen tube growth have been reported for the Nymphaeaceae (Orban and Bouharmont, 1995) and magnoliid taxa to include the Saururaceae (Pontieri and Sage, 1999) and Winteraceae (Sage et al., 1998; Sage and Sampson, 2003). In the cases where SI is present, self-recognition and rejection does not occur on the ventral carpel surface at the site of carpel closure. Instead, mechanisms of self-pollen recognition and rejection function either to prevent bicellular pollen growth at a dry stigma (Pontieri and Sage, 1999; Bernhardt et al., 2003; Hristova et al., 2005), a process traditionally viewed as uncommon (de Nettancourt, 1997; 2001; Sage et al., 2000), or inhibit embryo development (late-acting ovarian SI; Sage et al., 1998; Sage and Sampson, 2003).
The use of histochemistry and cryofixation techniques that optimize cellular retention of the transmitting tissue ECM have demonstrated that the esterase-positive pellicle and cuticle proper of the stigma overlay an ECM that is similar in structure to a cuticular layer that contains cutin, lipids, high and low methyl-esterified homogalacturonans, and arabinogalactan-proteins (AGPs) in Amborella (Thien et al., 2003), Chloranthus, Sarcandra (Hristova et al., 2005), and Kadsura (Lyew et al., 2007). Comparative quantitative and qualitative analyses of high and low methyl-esterified homogalacturonan and AGP distribution between the ECM of stigmatic epidermal cells and dorsal epidermal carpel cells not involved in pollen tube transmission in Chloranthus, Sarcandra (Hristova et al., 2005), and Kadsura (Lyew et al., 2007) indicate that the stigmatic ECM is structurally, histochemically, and immunochemically different from the ECM of dorsal epidermal carpel cells.
The presence of the esterase-positive pellicle and cuticle proper overlying a cuticular layer containing lipids, methyl-esterified homogalacturonans and AGPs presumably reflects an important role for these ECM constituents in the interactions between the pollen grain/pollen tube and the stigma, as well as pollen tube growth beyond the stigma of the basal grade angiosperms, as has been demonstrated in monocots and eudicots (Edlund et al., 2004; Swanson et al., 2004; Johnson and Lord, 2006; Hiscock and Allen, 2008). The pellicle is essential for compatible pollen–stigma interactions in monocots and eudicots (Heslop-Harrison and Heslop-Harrison, 1975). Esterases within the pellicle may include cutinases and pectin methyl esterases (Heslop-Harrison and Heslop-Harrison, 1975; Knox et al., 1976; Shaykh et al., 1977; Hiscock et al., 2002a; Swanson et al., 2005; Tung et al., 2005; Hiscock and Allen, 2008) that function either independently of, or in combination with, similar enzymes in the pollen ECM (Knox et al., 1976; Hiscock et al., 2002a; Suen et al., 2003) in remodeling the stigmatic ECM to aid pollen tube penetration therein. Additional cell wall modifying enzymes may be part of the stigmatic ECM exclusive of the pellicle (Bednarska et al., 2005; Swanson et al., 2005; Tung et al., 2005).
Lipids, which are present in the stigma ECM of both monocots and eudicots (Heslop-Harrison and Heslop-Harrison, 1982), may serve a common function during pollination (Hiscock and Allen, 2008). Lipids in wet stigmas have been demonstrated to be important in pollen hydration and pollen tube guidance by forming a water gradient between the hydrophobic and hydrophilic phases of the ECM (Wolters-Arts et al., 1998, 2002). Pollen grains and pollen tubes are presumed to respond to this gradient by hydrating and germinating directionally. Whether lipids in the ECM of dry stigmas are also instrumental in pollen hydration and germination remains to be demonstrated. Some species with dry stigmas appear to lack lipids, and it has been posited that, in these species, the role for lipids has been transferred from the stigma to the pollen grain ECM with the evolution of a dry stigma from a wet stigma (Dickinson, 1995; Lush et al., 2000; Hiscock and Allen, 2008).
Methyl-esterified homogalacturonans and AGPs, which are well known to be present within the ECM of the growing pollen tube, stigma, and style of monocots and eudicots, have numerous biological functions (Heslop-Harrison and Heslop-Harrison, 1982; Li et al., 1995; Juah and Lord, 1996; Lennon et al., 1998; Hasegawa et al., 2000; Lennon and Lord, 2000; Mollet et al., 2000, 2002; Park et al., 2000; Wu et al., 2000; Lenartowska et al., 2001; Showalter, 2001; Khosravi et al., 2003; Coimbra and Duarte, 2003; Bednarska et al., 2005). Methyl-esterified homogalacturonans and AGPs are primary components of the primary wall of the pollen tube (Geitmann and Steer, 2006). Synthesis and secretion of AGPs and high methyl-esterified homogalacturonans occur at the growing tip of the pollen tube; subsequent de-esterification results in the presence of low methyl-esterified homogalacturonans within the pollen tube wall (Bosch et al., 2005; Geitmann and Steer, 2006). Methyl-esterified homogalacturonans and AGPs within the pollen wall are presumed to provide mechanical stability during siphonogamous pollen tube growth (Geitmann and Steer, 2006).
Low methyl-esterified homogalacturonans and AGPs within the pollen tube wall and transmitting tissue ECM may also operate in pollen tube adhesion and guidance of pollen tubes within stylar transmitting tissue (Li et al., 1995; Roy et al., 1998; Mollet et al., 2000, 2002; Park et al., 2000; Wu et al., 2000; Lenartowska et al., 2001; Coimbra and Duarte, 2003; Kim et al., 2006). Adhesive molecules in transmitting tissue are posited to function in pollen tube guidance by keeping pollen tubes on a prescribed path (Johnson and Lord, 2006). Along these lines, transmitting tissue epitopes recognized by JIM5, a monoclonal antibody that binds to low methyl-esterified homogalacturonans, have been demonstrated to interact with a cysteine-rich adhesion (SCA) protein in Lilium longiflorum to participate in adhesion between stylar transmitting cells and pollen tubes (Mollet et al., 2000). The cysteine-rich adhesion protein of L. longiflorum belongs to the family of lipid transfer proteins (LTPs; Mollet et al., 2000). Other proteins within this family have been shown to have an important role in pollen tube growth at the stigma by having expansin-like cell wall loosening characteristics (Nieuwland et al., 2005), leading Hiscock and Allen (2008) to conclude that LTPs may also be one broad category of compounds involved in pollen tube growth in angiosperms. LTPs have also been implicated in cuticle synthesis (Samuels et al., 2008). Recently, SCA has been demonstrated to interact with ubiquitin in pollen tube adhesion in lily (Kim et al., 2006); both molecules are endocytosed at the growing pollen tip. At present, the functional significance of this process remains unclear. Immunolocalization studies indicated the presence of epitopes recognized by anti-SCA within the stigmatic ECM where pollen tubes grow intercellularly into the "solid" transmitting tissue of the basal taxon, Chloranthaceae (Hristova et al., 2005), providing limited support for the view that LTPs may also be universally involved in pollen tube growth in angiosperms.
Numerous other functions have been assigned to methyl-esterified homogalacturonans and AGPs within the ECM of the pollen grain, pollen tube, and transmitting tissue. Low methyl-esterified homogalacturonans may operate as calcium reservoirs to support pollen tube tip growth (Lenartowska et al., 2001; Bednarska et al., 2005). Methyl-esterified homogalacturonans and AGPs are also presumed to play a role in hydration and stabilization of the transmitting tissue ECM architecture (Carpita and Gibeaut, 1993) and in pollen tube nutrition and cell signaling (Cheung et al., 1995; Lind et al., 1996; Youl et al., 1998). The diversity in functions attributed to methyl-esterified homogalacturonans and AGPs during pollen–carpel interactions in species from a wide range of taxonomic affinities suggests that, like lipids, methyl-esterified homogalacturonans and AGPs may be universally important in their support of angiosperm pollen tube growth.
The view that the wet stigma predates the dry stigma during angiosperm history (Dickinson, 1995, p. 9), has led to the question "...in the evolution of dry stigmas, at what stage did the pollen begin to secrete cutinase and when did a new generation of pollen coatings develop, capable of adhesion, transfer of water, and playing a role in the SI system?" The pollen wall of relictual-basal angiosperms contains a pollen coating (Furness and Rudall, 2001; Koehl, 2002; Thien et al., 2003). Histochemical studies indicate that this coating is esterase-positive and contains homogalacturonans and lipids in Amborella (Thien et al., 2003) and Illicium (Koehl, 2002). These data, in combination with the demonstration of an esterase-positive dry stigma in other basal-relictual angiosperms, suggests that ECM compounds involved in adhesion, transfer of water, and penetrative siphonogamous pollen tube growth may have been present within the pollen coating and stigmatic ECM during the early history of cell-to-cell interactions between pollen grains/pollen tubes and stigmatic cells. Similarly, the presence of stigmatic SI in combination with bicellular pollen grains in basal-relictual angiosperms that function either to prevent pollen grain hydration (Pontieri and Sage, 1999; Pontieri, 2004; Hristova et al., 2005) or pollen tube penetration of stigmatic tissue (Bernhardt et al., 2003) provides the intriguing possibility that stigmatic SI evolved much earlier in angiosperm history than has been predicted (Dickinson, 1995; Weller et al., 1995).
The purpose of the current study is to continue to characterize the architectural features of transmitting tissue ECM and pollen tube growth in extant representatives of basal-relictual angiosperm lineages to test hypotheses regarding the early evolution and function of angiosperm transmitting tissue. One objective is to use histochemistry and high resolution imaging of cryofixed transmitting tissue to assess whether the ECM of the stigma of Illicium floridanum (Illiciaceae), a member of the ANITA grade (Qiu et al., 1999), and Acorus americanus of the family Acoraceae, which is sister to the rest of the monocots (Tamura et al., 2004; Saarela et al., 2007), is similar to a wet or dry stigma and whether the transmitting tissue ECM beyond the stigma up to the micropyle in Illicium and Acorus, as well as in Amborella and Trimenia moorei is freely flowing.
A second goal is to characterize whether high and low methyl-esterified homogalacturonans and AGPs are present within the transmitting tissue ECM at the time of stigma receptivity for Amborella, Illicium, Trimenia, and Acorus. For comparative purposes, we assess the structural and immunochemical architecture of dorsal epidermal cells not involved in pollen tube growth within these species. Given the phylogenetic position of Amborella, we also provide a developmental analysis of the structure and immunochemical make-up of the stigmatic and dorsal epidermal ECM with respect to high and low methyl-esterified homogalacturonans and AGPs. Combined, these data will determine whether compounds deemed to be universally important in pollen–carpel interactions are present within the transmitting tissue and whether the transmitting tissue ECM is immunochemically distinct from the ECM of epidermal cells not involved in pollen tube growth.
The third objective is to document the structural and immunochemical features of high and low methyl-esterified homogalacturonans and AGPs within the microenvironment between the ECM of the pollen grain and stigma and that of the pollen tube and stigma in Amborella and Illicium. The following questions will be addressed: (1) Are the initial interactions between the pollen grain and the stigma comparable to those of other species with a dry stigma following a compatible pollination with respect to pollen adhesion, hydration, and remodeling of the ECM? (2) Do the cellular interactions between the pollen grain/pollen tube and transmitting tissue ECM resemble those of monocots and eudicots with respect to methyl-esterified homogalacturonans and AGPs, thereby implicating these ECM components in pollen adhesion, hydration, germination, and pollen tube growth?
The fourth objective is to use the observations on the architecture of the transmitting tissue ECM to test the hypothesis that the ancestral transmitting tissue ECM is homologous with the pollination droplet of gymnosperms. A prediction of this hypothesis is that a freely flowing stigmatic ECM was ancestral to a dry stigmatic ECM in the angiosperms as a whole. We test this prediction by reconstructing the character phylogeny of freely flowing stigmatic ECM in a sample of species that represents most early-diverging angiosperm lineages to include those examined in the current study.
The final goal of the current study is to place the combined results of this research in the context of data from previous studies addressing cuticle development in angiosperms to generate a refined hypothesis on the evolution of transmitting tissue—a major floral innovation.
MATERIALS AND METHODS
Plant material
Samples were obtained from a minimum of three plants each of Amborella trichopoda, Illicium floridanum, Trimenia moorei, and Acorus americanus. Floral tissue was harvested from Amborella plants growing in New Caledonia at Col d’Amieu and Plateau de Dogny as described by Thien et al. (2003) . Floral tissue from Trimenia was acquired from plants collected as described by Bernhardt et al. (2003) and subsequently cultivated in pots in a shadehouse at the Royal Botanical Gardens, Sydney. Flowers of Illicium were obtained from native populations as described by Koehl (2002) and plants were grown in controlled growth rooms at the University of Toronto greenhouse facility. The Doremus Wholesale Nursery (Warren, Texas) and the Banting Family Nursery Farms (Bridge City, Louisiana) were the original sources for Illicium that was cultivated at the University of Toronto. Plants of Acorus were collected at High Park, Toronto, Ontario (43°38'38.62''N, 79°28'04.91''W) and were also grown in the University of Toronto greenhouse facility. Controlled pollinations on Amborella were performed in New Caledonia as described by Thien et al. (2003) . Controlled pollinations on Illicium were conducted on plants growing in either native populations (Koehl et al., 2004) or at the University of Toronto. Pollinations on Acorus were completed at the University of Toronto.
ECM structure and composition of transmitting tissue
The ECM architecture and composition of the transmitting tissue in Amborella, Illicium, Trimenia, and Acorus before and after pollination were characterized by preparing carpels for scanning electron microscopy (SEM) and transmission electron microscopy following cryofixation as described by Lam et al. (2001) and Thien et al. (2003) . Unpollinated gynoecia (N 
7 flowers/plant) were sampled at the time of stigma receptivity (Amborella, Illicium, Trimenia, and Acorus). Unpollinated gynoecia (N 7 flowers/plant) of Amborella were also sampled from 1- and 2-mm floral buds. Pollinated gynoecia of Amborella, Illicium, and Acorus were harvested 1 and 6 h after pollination (N 5 flowers/plant/sample time).
Cryofixed unpollinated and pollinated gynoecia were also used to determine the presence and spatial distribution of methyl-esterified homogalacturonans and AGPs in the ECM of transmitting tissue, pollen grains, pollen tubes, and dorsal carpel cells not involved in pollen tube growth. Homogalacturonans and AGPs were detected using monoclonal antibodies that recognize epitopes of highly methyl-esterified homogalacturonans (JIM7), low methyl-esterified homogalacturonans (JIM5), and AGPs (JIM13) as previously described (Hristova et al., 2005; Lyew et al., 2007). The development of transmitting tissue ECM in Amborella was characterized by quantifying immunolocalization events of all JIMs on an area basis as described by Hristova et al. (2005) . Density values for each antibody were determined at the outermost surface of the stigma at the tip of uniseriate, multicellular stigmatic papillae, between the rows of uniseriate, multicellular stigmatic papillae, and between the cells composing the individual uniseriate, multicellular papillae. The same density values for each antibody were also determined at the surface of dorsal epidermal carpel cells, between dorsal epidermal carpel cells, and at the base of dorsal epidermal carpel cells to determine if stigmatic epidermal cells were distinct from other carpel epidermal cells. First, all density values at each epidermal location for each antibody/carpel were contrasted using one-way ANOVA (Sigma Stat 2.03, Chicago, Illinois, USA). No differences were observed between carpels; therefore, all values per carpel per antibody were pooled for each location at each respective epidermal position. Density values per antibody were then contrasted.
To determine whether a cuticle and pellicle were present in the receptive stigmas of Illicium and Acorus, fresh, whole flowers were also examined using the following stains: (1) 0.01% auramine O in 0.05 M Tris/HCL buffer at pH 7.2 observed under UV light to detect cutin and lipids (Heslop-Harrison and Shivanna, 1977), and (2) indoxyl acetate in 0.1 M tris/HCl buffer at pH 7.0 in 0.1 M potassium ferrocyanide and potassium ferricyanide to detect nonspecific esterase activity in the stigma pellicle (De Jong et al., 1967). Controls for nonspecific esterase activity were run by omitting the substrate. Controls for remaining histochemical reagents were run by omitting the stain. Confirmation of the distribution of transmitting tissue ECM requires an analysis of spatial patterns of pollen tube growth. The pollen tube pathway has been previously determined for all of the species studied here (Williams et al., 1993; Thien et al., 2003; Bernhardt et al., 2003; Koehl et al., 2004) with the exception of Acorus. Therefore, stigmas from 25 plants of Acorus were cross- (N = 25 flowers/plant) and self-pollinated (N = 25 flowers/plant) at stigmatic receptivity and prepared for aniline blue fluorescence microscopy 24 and 48 h after pollination as described by Martin (1959) .
Reconstruction of character phylogeny
Data from the current study and published data for presence or absence of a freely flowing stigmatic ECM in representatives of Amborellales (present study), Nymphaeales (Heslop-Harrison and Shivanna, 1977; Schneider and Chaney, 1981), Austrobaileyales (present study; Lyew et al., 2007), Chloranthales (Hristova et al., 2005), Magnoliidae (Sage and Sampson, 2003), and selected monocotyledons (present study; Heslop-Harrison and Shivanna, 1977; Ciampolini et al., 2001; Sage et al., 2001) and eudicotyledons (Heslop-Harrison and Shivanna, 1977; Bednarska, 1991; Elleman et al., 1992) were tabulated and traced onto the relevant branches of the angiosperm cladograms illustrated in Stevens (2001) using the parsimony and maximum likelihood options of Mesquite version 2.0 (Maddison and Maddison, 2007). The evolutionary models implemented were unweighted, unordered parsimony and the Markov k-state 1-parameter model of Lewis (2001) for maximum likelihood.
RESULTS
Transmitting tissue ECM: Development, structure, and composition in unpollinated gynoecia
Amborella
The mature stigma is composed of uniseriate, multicellular papillae (Endress and Igersheim, 2000a, b). These stigmatic cells are already apparent in gynoecia from 1-mm floral buds (Fig. 1A, inset). A cuticle overlays the primary wall of the surface of stigmatic epidermal cells of gynoecia from 1- and 2-mm floral buds and flowers at anthesis (Fig. 1A–M). This cuticle remains continuous even though periclinal divisions result in an increase in the length of individual files of uniseriate, multicellular papillae throughout the developing stigma. The cuticle is composed initially of a cuticle proper (Fig. 1D) and subsequently a cuticle proper and reticulate cuticular layer (Fig. 1F, 1J–M). The reticulate network of the cuticular layer extends from the primary wall to the surface of the cuticle proper (Fig. 1F, 1J, 1K–M). The cuticle proper and cuticular layer are unevenly thickened with outgrowths (Fig. 1G–I) defined previously by Heslop-Harrison and Heslop-Harrison (1982) as cutinized bosses. Although the cuticle remains continuous throughout stigma development, the cuticle proper associated with the bosses occasionally appears discontinuous (Fig. 1K). Lipidic inclusions are present within the bosses (Fig. 1K) and throughout the cuticular layer.
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The transmitting tissue ECM at the ventral surface of the carpel beyond the stigma and up to the micropyle is composed almost entirely of a primary wall covered by a cuticle proper at anthesis (Fig. 1S–V). On occasion, an unevenly expanded cuticular layer is present (Fig. 1U). The micropyle contains a prominent ECM beyond the primary wall of the inner integument and nucellus (Fig. 1W, 1 X). This ECM also consists of a cuticle proper and a cuticular layer with prominent lipidic occlusions (Fig. 1 X).
The ECM of transmitting tissue and dorsal carpel epidermal cells not involved in pollen tube growth contains epitopes that are recognized by monoclonal antibodies JIM 7, 5, and 13. The primary wall of the surface of the stigma, as well as the reticulate regions of the cuticle, contains epitopes recognized by JIM7 (Fig. 1E, 1F, 1J, 1K), JIM5 (Fig. 1L), and JIM13 (Fig. 1M). JIM7 also localizes at the expanded cuticular layer between individual uniseriate, mulitcellular stigmatic papillae and the primary wall of cells making up individual uniseriate stigmatic papillae (Fig. 1P), the ventral carpel epidermis (Fig. 1U), and nucellar and inner integument epidermis (Fig. 1 X). Epitopes recognized by JIM5 (Fig. 1Q) and JIM13 (Fig. 1R) are also present in the expanded cuticular matrix between individual files of uniseriate stigmatic papillae but are rarely present in the primary wall of the uniseriate multicellular stigmatic papillae, ventral epidermal carpel cells, and nucellar and inner integument epidermal cells. When present in the ventral epidermal carpel cells (Fig. 1V) and nucellar and inner integument epidermal cells, JIM13 epitopes localize at the plasma membrane. Epitopes recognized by JIM 7 (Fig. 1D, inset) and JIM5 within the dorsal epidermal cells localize at the primary wall, whereas those recognized by JIM13 are present at the plasma membrane.
Epitopes recognized by JIM7 display a higher density than those recognized by JIM5 and 13 at all ECM locations in stigmas from 1- and 2-mm floral buds and the dorsal epidermis of the carpel at all developmental stages (Fig. 2A–C). The density of epitopes recognized by JIM7 remain relatively unchanged over time except for a significant decline within the expanded cuticular layer between uniseriate, multicellular stigmatic papillae at anthesis which corresponds to an increase in density of epitopes recognized by JIM5 and 13 (Fig. 2A–C). The density of epitopes recognized by JIM7, 5, and 13 in the ECM of the dorsal carpel epidermal cells remain relatively constant throughout gynoecial development and are in most cases significantly less than that in the stigmatic ECM (Fig. 2A–C).
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; Fig. 3A), which is composed of unicellular papillate cells (Robertson and Tucker, 1979; Williams et al., 1993; Fig. 3B, 3C). The ECM of stigmatic cells is composed of a primary wall overlaid by a cuticle proper and a reticulate cuticular layer, which contains lipidic inclusions and frequently forms isolated cuticular bosses (Fig. 3B–H). The reticulate network of the cuticular layer extends from the primary wall to the surface of the cuticle proper (Fig. 3D–H). The cuticle is positive for nonspecific esterase activity (Fig. 3C, inset) and auramine O (Fig. 3D, inset). The dorsal carpel epidermal cell ECM is composed of a uniformly thickened rugose cuticle and a primary wall (Fig. 3E inset, 3F inset).
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The ECM of transmitting tissue contains epitopes that are recognized by JIM7, 5, and 13. The primary wall of the stigma surface contains epitopes recognized by JIM7 (Fig. 3E), whereas epitopes recognized by JIM5 (Fig. 3F, 3G) and 13 (Fig. 3H) are present in the primary wall and reticulate regions of the cuticular layer and cuticle proper. JIM7 localizes at the primary wall of the ventral carpel epidermis (Fig. 3L), and JIM13 localizes at the cuticular layer of the ventral carpel epidermis (Fig. 3M). Epitopes recognized by JIM7, 5 (Fig. 3O), and 13 are present in the micropylar ECM. Although JIM7 and 5 localize at the primary wall of dorsal epidermal cells (Fig. 3E inset, 3F inset), epitopes recognized by JIM13 are absent.
Trimenia
The mature stigma has multicellular protrusions (Endress and Sampson, 1983; Fig. 4A–C). The surface ECM of stigmatic cells is composed of a primary wall overlaid by a thin cuticle (Fig. 4D). In contrast, the primary wall of dorsal epidermal carpel cells is covered with a thick rugose cuticle (Fig. 4D inset, 4E inset). A continuous expanded ECM extends from the lateral walls of stigmatic cells (Fig. 4F) basipetally to the ground tissue and the ventral carpel surface at the site of carpel closure (Bernhardt et al., 2003; Fig. 4G). The ECM of adjacent ventral carpel surfaces is fused (Fig. 4G, 4H). An ECM that extends beyond the primary wall of the nucellar and inner integument epidermis is present in the micropyle (Fig. 4I).
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Acorus
Controlled hand cross- and self-pollinations of A. americanus that were conducted to determine the spatial distribution of transmitting tissue resulted in only one successful cross-pollination. The unsuccessful cross- and self-pollinations are due to a lack of pollen hydration at the stigmatic surface (Fig. 5A). The successful cross-pollination results in germination of pollen on the stigma and subsequent growth of pollen tubes along the ventral sides of the carpel at the site of carpel closure into the ovary (Fig. 5B, 5C). From these observations, we concluded that the site of carpel closure beyond the stigma is the site of transmitting tissue leading to the ovary locule.
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; Fig. 6A) that are positive for nonspecific esterase activity (Fig. 6A, inset) and auramine O (Fig. 6B, inset). A cuticle composed of a cuticle proper and reticulate cuticular layer overlays the primary wall of the surface of stigmatic epidermal cells (Fig. 6B–E). Unicellular papillate epidermal cells are also present on the ventral surface of the carpel at the site of carpel closure (Buzgo and Endress, 2000), and an expanded ECM is present beyond the primary walls of these cells (Fig. 6F, 6H, 6I). This ECM is continuous with an abundant ovarian ECM (Fig. 6G, 6J, 6K). The ovarian ECM, which is derived from epidermal cells (Rudall et al., 1998), has a well-defined boundary (Fig. 6K) and is structurally similar to a reticulate cuticular layer (Fig. 6J, 6K) that has no obvious cuticle proper. The ECM of the dorsal carpel epidermal cells is composed of a uniformly thickened smooth cuticle and a primary wall (Fig. 6D, inset). Lipidic occlusions are apparent in the ECM of the stigma and the site of carpel closure (Fig. 6D, 6H, 6I, inset).
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Structural and immunological features of the microenvironment between the ECM of the pollen grain/pollen tube and stigmatic transmitting tissue
Amborella
The ECM between the stigma and pollen grain/germinating pollen tube are contiguous and often indistinguishable from one another (Fig. 7A–R). Interactions between the pollen grain and stigma ECM are limited to the point of contact between the two cells (Fig. 7A–G). The microenvironment within the adhesion zone at the stigma surface is composed of both pollen and stigma ECM and contains epitopes recognized by JIM7 (Fig. 7B, 7D), 5 (Fig. 7G), and to a lesser degree, JIM13 that originated from the stigmatic and pollen ECM (Fig. 7C, 7F). The pollen ECM also contains a lipidic component (Fig. 7B, 7C, 7F), which, like the JIM7- (Fig. 7B, 7C), 5- (Fig. 7F, 7G), and 13-positive ECM, is associated with the interstices and periphery of the exine. The foot layer of the exine as well as the intine also contain epitopes recognized by JIM7 (Fig. 7B, 7C), 5 (Fig. 7F, 7G), and 13.
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The ECM at the apex of pollen tubes is ill defined and contiguous with a distinctly demarcated subapical wall (Fig. 7E, 7H–J, 7M). JIMs 7, 5, and 13 localize at the well-defined subapical wall (Fig. 7E, 7H–J). Callose is also present in the pollen tube wall (Fig. 7H, 7L, 7N, 7Q, 7R) and adjacent to the intine of the hydrated pollen grain (Fig. 7A). Epitopes recognized by JIM7 (Fig. 7E) and 13 (Fig. 7J) are present in the ECM and plasma membrane of the amorphous apical tip, respectively. The growing pollen tube is composed of an apical cytoplasmic region that contains mitochondria and vesicles that were immunopositive with JIM7 (Fig. 7E, 7M) and 13 (Fig. 7J, 7K). A subapical cytoplasmic zone contains mitochondria, plastids, rough endoplasmic reticulum (Fig. 7N–P), and Golgi-derived vesicles that are immunopositive with JIM7 (Fig. 7O) and 13 (Fig. 7P).
Illicium
The ECM between the stigma and pollen grain and between the stigma and germinating pollen tube form an adhesion zone that is frequently homogeneous at the point of contact (Fig. 8A–R). The microenvironment within the adhesion zone is composed of both pollen and stigma ECM and contains epitopes recognized by JIM7 (Fig. 8B, 8C, 8M), 5 (Fig. 8E, 8F, 8N, 8O), and 13 (Fig. 8I–K, 8P–R). Lipidic components are also present within the pollen grain ECM (Fig. 8B–K), which, like the JIM7- (Fig. 8B–D), 5- (Fig. 8E–G), and 13-positive (Fig. 8H–K) pollen ECM, is associated with the interstices and periphery of the exine. The foot layer of the exine, as well as the intine, also contains epitopes recognized by JIM7 (Fig. 8B–D), 5 (Fig. 8E), and 13 (Fig. 8H-K).
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The apex of the pollen tube wall is immunopositive with JIM7 (Fig. 8M) and 13 (Fig. 8P, 8Q), whereas the pollen tube wall behind the apex contains epitopes recognized by JIM5 (Fig. 8N, 8O) and 13 (Fig. 8R). Callose is also present in the pollen tube wall behind the apex (Fig. 8N, 8O). The growing pollen tube is composed of an apical zone that contains mitochondria and vesicles that are immunopositive with JIM7 (Fig. 8M) and 13 (Fig. 8P, 8Q). A subapical zone of growing pollen tubes contains mitochondria, plastids, rough endoplasmic reticulum, and Golgi-derived vesicles that are immunopositive with JIM7 and 13 (Fig. 8R).
Reconstruction of character phylogeny
Data for the presence or absence of a freely flowing stigmatic ECM were tabulated for 27 species of angiosperms representing the taxa Amborellales (1 sp.), Nymphaeales (4 spp.), Austrobaileyales (3 spp.), Chloranthales (2 spp.), Magnoliidae (3 spp.), monocotyledons (8 spp.) and eudicotyledons (6 spp.). This sample of taxa includes descendants of 13 of the 16 basalmost nodes in the angiosperm cladogram of Stevens (2001) . The most parsimonious reconstruction of the character phylogeny unequivocally resolves dry stigma as ancestral for the angiosperms as a whole, as well as for all of the clades listed (Fig. 9). Freely flowing stigmatic ECM is resolved as originating independently at least four times in the Nymphaeaceae, Winteraceae, Asparagales, and Solanaceae (Fig. 9). Forcing the sampled occurrences of freely flowing stigmatic ECM to be homologous and ancestral would require 13 independent origins of dry stigmas. The maximum likelihood reconstruction of ancestral states (result not shown) was essentially the same as the parsimony result, resolving dry stigma as ancestral to the angiosperms as a whole (with a proportional likelihood of 0.98) and to most internal internodes on the tree.
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Is transmitting tissue ECM freely flowing in Amborella, Illicium, Trimenia, and Acorus?
Data from the current study clearly demonstrate that the stigmatic ECM of Illicium and Acorus is of the dry-type as indicated by the presence of an esterase-positive cuticle overlying a primary wall at receptivity. The cuticle of the unicellular stigmatic cells is composed of a cuticle proper and an expanded cuticular layer in each species. As well, observations on development of the dry-type stigma in Amborella illustrate that the cuticle of each uniseriate, multicellular papilla is also composed of a cuticle proper and expanded cuticular layer. In Amborella, the cuticular matrices from adjacent papillae become fused during development. This fused stigmatic matrix where pollen tubes will grow in Amborella (Thien et al., 2003) structurally resembles a "solid style" in other angiosperm species (Tilton and Horner, 1980; Cresti et al., 1992) that is composed of long files of cells, which are, as noted here for Amborella, connected by plasmodesmata and surrounded by an expanded ECM. This cellular architecture has been proposed to provide important topographic guidance for pollen tubes (Heslop-Harrison et al., 1985; Lush et al., 2000), likely functioning in tandem with other pollen tube guidance mechanisms (Lush et al., 2000).
Collectively, a dry stigma has now been demonstrated to be present in seven basal-relictual angiosperm taxa to include Amborellaceae (present study; Thien et al., 2003; Hristova et al., 2005), Illiciaceae (present study; Koehl, 2002), Schisandraceae (Lyew et al., 2007), Trimeniaceae (Bernhardt et al., 2003), Chloranthaceae (Hristova et al., 2005), Saururaceae (Pontieri and Sage, 1999; Pontieri, 2004) and Acoraceae (present study). Wind pollination occurs in Brasenia (Osborn and Schneider, 1988; Endress, 2005) indicating that a dry stigma may also develop within the Cabombaceae. And, because most species within the Hydatellaceae flower under the water (Rudall et al., 2007), it is unlikely that a stigma with a freely flowing secretion is present. Species within the Nymphaeaceae have either wet or dry stigmas (Heslop-Harrison and Shivanna, 1977; Schneider and Chaney, 1981).
The transmitting tissue ECM extending beyond the stigma up to the embryo sac in carpels of Amborella, Illlicium, Trimenia, and Acorus is also not freely flowing. Unexpectedly, the ECM of the ventral epidermis below the stigma up to the ovary locule where pollen tubes grow after exiting the stigma in Amborella (Thien et al., 2003) and Illicium (Williams et al., 1993; Koehl et al., 2004) bears limited ultrastructural resemblance to the stigma ECM. This ventral epidermis develops a comparatively thin matrix composed primarily of a smooth cuticle proper beyond the primary wall. An unevenly expanded cuticular layer overlaid by the cuticle proper is occasionally present. Even though the ECM at this ventral carpel surface has been proposed to function in both pollen tube growth and sealing of the carpel (Endress and Igersheim, 2000a), evidence from the current study indicates that it only serves the former function in these two species because the ECM of the adjacent ventral surfaces remain unfused. The fused cuticular stigmatic ECM operates solely in sealing of the carpel in Amborella.
The nonfreely flowing ECM in Trimenia and Acorus at the ventral surfaces of the carpel beyond the stigma and up to the ovary locule is notably different in structure and function from that of Amborella and Illicium. In comparison to Amborella and Illicium, this ECM in Trimenia and Acorus is abundant beyond the primary wall and functions in carpel closure. This copious matrix operates in sealing the carpel presumably due to the abundance of low methyl-esterified homogalacturonans in this region. Low methyl-esterified homogalacturonans can aggregate via calcium bridges and form gels that are essential for cellular adhesion (Goldberg et al., 1996; Willats et al., 2001; Iwai et al., 2002). And, while the current study provides some evidence that this ECM also supports compatible pollen tube growth following stigmatic germination in Acorus, a previous study (Bernhardt et al., 2003) indicates that this ECM does not operate in compatible pollen tube growth in Trimenia. Rather, Trimenia pollen tubes grow within an ECM that is presumably derived from ground tissue, which extends up to the stigmatic ECM and is contiguous with the ventral epidermal ECM (present study; Bernhardt et al., 2003).
Are methyl-esterified homogalacturonans and AGPs present within the transmitting tissue ECM of Amborella, Illicium, Trimenia, and Acorus?
The current study clearly demonstrates that methyl-esterified homogalacturonans and AGPs, two classes of compounds deemed important for pollen–carpel interactions in monocots and eudicots, are present in the transmitting tissue ECM of all species. And, even though these molecules are also present within the ECM of carpel epidermal cells not involved in pollen tube growth, there are outstanding differences in the spatial array of these molecules between the two cell types that are likely important for pollen grain adhesion, hydration, and germination at the stigmatic ECM. Homogalacturonans are located within a reticulate network of both the cuticle in stigmatic and dorsal epidermal cells. However, the retiform of homogalacturonans extends to the surface of the cuticle of stigmatic cells only. These results are noteworthy because even though the stigmatic cuticle itself is, in each taxon, continuous over the stigma, the surface of the stigma is not rendered impermeable by a continuous layer of lipophilic molecules that make up the cuticle proper. Rather, it contains hydrophilic molecules that impart porosity as well as rigidity to the plant cell ECM (Carpita and Gibeaut, 1993; Baron-Epel et al., 1988; Cosgrove, 2005). AGPs are additionally present within the cuticular layer of Amborella and Illicium stigmatic cells where they may also increase ECM porosity (Seifert and Roberts, 2007). This spatial distribution of homogalacturonans observed in the stigmatic cuticle of Amborella, Illicium, Trimenia and Acorus has also been documented to occur in Kadsura longipedunculata (Schisandraceae) and two species within the Chloranthaceae (Hristova et al., 2005). We hypothesize that the hydrophilic molecular network provided by homogalacturonans and AGPs in the stigmatic ECM of these basal angiosperms enhances cuticle permeability and acts as an avenue for the movement of water and other molecules functioning in cellular interactions between the stigma and pollen.
The stigmatic ECM of Amborella undergoes dramatic changes in homogalacturonan and AGP composition during the course of maturation. First, cuticular blisters, also described as bosses (Heslop-Harrison and Heslop-Harrison, 1982), which are absent early in stigma development, are prominent at the stigmatic surface where pollen grains will adhere at anthesis. These bosses form within the cuticular layer and contain high and low methyl-esterified homogalacturonans, AGPs as well as electron-translucent regions interpreted to be composed of lipid-based molecules (Hristova et al., 2005; Lyew et al., 2007). These bosses do not form within the ECM of dorsal carpel epidermal cells that are not involved in pollen tube growth. Similar cuticular bosses are also present in other taxa with dry stigmas, such as two species of Chloranthaceae (Hristova et al., 2005), the magnoliid Saururaceae (Pontieri and Sage, 1999; Pontieri, 2004), and the eudicot Brassicaceae (Elleman and Dickinson, 1994, see also Heslop-Harrison and Heslop-Harrison, 1982). Second, the ECM at the surface of the Amborella stigmatic cells has a two-fold increase in the density of low methyl-esterified homogalacturonans at anthesis relative to earlier developmental stages, indicating the activity of de-esterification enzymes (Goldberg et al., 1996). Third, in comparison to the unchanging ECM between cells that make up an individual stigmatic papilla, the fused cuticular ECM between the stigmatic papillae, where pollen tubes will grow, enlarges dramatically at anthesis in concert with an approximately 30-fold increase in density of low methyl-esterified homogalacturonans and 5-fold increase in density of AGPs recognized by JIM13. Because anthesis is the time at which Amborella stigmas are receptive (Thien et al., 2003), the site-specific changes in molecules deemed important for pollen grain adhesion, hydration, germination, and pollen tube growth presumably operate to prepare the stigma ECM for these processes.
In addition to documenting the presence of high and low methyl-esterified homogalacturonans and AGPs in the transmitting tissue of the carpel, the present study illustrates that these important molecules also occur within a prominent ECM of the micropyle in Amborella, Illicium, and Trimenia. In general, angiosperm pollen tubes change 90° in their direction of growth when entering the micropyle, and the micropylar ECM is hypothsized to provide guidance for pollen tubes at this junction (Johnson and Lord, 2006). Micropylar exudates have been identified in a number of diverse angiosperms (Sage et al., 1994). The fine structure of integumentary and nucellar cells of Ornithogalum led Tilton (1980) to conclude that cells lining the micropyle are responsible for secretion of the micropylar contents. In addition, angiosperm micropylar exudates may also arise from degenerated nucellar cells (Chao, 1971; Sage and Williams, 1993; Sage et al., 1994) or the embryo sac itself (Sage and Williams, 1993; Sage et al., 1994). ECM components within the nucellus (Márton et al., 2005) and synergids (Kristóf et al., 1999) have also been documented to operate in guiding pollen tubes into the embryo sac. The micropylar ECM in Amborella, Illicium, and Trimenia is composed of the homogalacuronan-rich cuticular layer of nondegenerated nucellar and integumentary cells. And, while the micropylar ECM in Amborella is also composed of a cuticle proper, a cuticle proper was not observed in Illicium and Trimenia. The lack of a cuticle proper in Illicium and Trimenia may be due to the rapid expansion of the thickened cuticular layer. Alternatively, a cuticle proper may be present but not visible with the transmission electron microscope (Jeffree, 2006). A predominant ECM is also present in the micropyle of Acorus. However, this ECM appears to be part of the copious ECM that arises from ovarian trichomes (present study; Rudall et al., 1998). The ECM of the epidermal cells of the ovary locule in Amborella, Illicium, and Trimenia is composed only of the primary wall and a thin cuticle proper. The presence of a prominent micropylar ECM that is composed of homogalacturonans and AGPs is a novel observation for angiosperms.
Cell-to-cell interactions between the pollen grain/pollen tube and transmitting tissue in Amborella, Illicium, and Acorus
Results from the present study demonstrate that the initial interactions between the pollen grain and the stigma ECM of Amborella and Illicium are comparable to those of other taxa with a dry stigma following a compatible pollination. Pollination in Amborella and Illicium results in the typical well-defined adhesive zone that is limited to the point of contact between the pollen grain and the dry stigma (Elleman et al., 1988, 1992; Pontieri and Sage, 1999; Sage et al., 2001; Hiscock et al., 2002b; Pontieri, 2004). And, while the adhesion zone in Amborella and Illicium is composed of both the stigmatic and pollen grain ECM, as commonly noted in the dry-type system (Elleman et al., 1988, 1992; Pontieri and Sage, 1999; Sage et al., 2001; Hiscock et al., 2002b; Pontieri, 2004), our observations provide new information on the pollen ECM that has significant implications for cell-to-cell interactions within the adhesion microenvironment. As observed for the stigma ECM, the pollen ECM contains high and low methyl-esterified homogalacturonans, AGPs, and lipids. The high and low methyl-esterified homogalacturonans and AGPs are tightly associated with the pollen foot layer, tectum, and columella. As well, these molecules form a boundary at the periphery of lipid inclusions located within the interstices and periphery of the exine. This homogalacturonan, AGP and lipid pollen ECM corresponds to the carbohydrate and lipid containing exinic outer layer in Brassica described by Gaude and Dumas (1984) . Adhesion of the pollen grain is hypothesized to involve interactions between esterases and other compounds within the pellicle and exinic outer layer (Hiscock and Allen, 2008). Consistent with this idea is the observation that a lipophilic, carbohydrate-based molecule that is tightly associated with the exine is involved in pollen grain adhesion in Arabidopsis (Zinkl et al., 1999).
We hypothesize that low methyl-esterified homogalacturonans present in the exinic outer layer and stigma ECM facilitate adhesion of pollen grains to the stigma in Amborella and Illicium. Adhesion may result from bridges formed between low methyl-esterified homogalacturonans and other ECM molecules (Mollet et al., 2000) or directly from interactions between low methyl-esterified homogalacturonans. As noted previously, low methyl-esterified homogalacturonans can aggregate via calcium bridges and form gels that are essential for cellular adhesion (Goldberg et al., 1996; Willats et al., 2001; Iwai et al., 2002). The pollen wall and stigma of Illicium and Amborella is esterase positive, as is the adhesion zone (Koehl, 2002; T. L. Sage, personal observations). Although a positive esterase response may reflect the presence of cutinase, it can also be indicative of pectin methyl esterase (Hiscock and Allen, 2008) that would effectively increase the number of low methyl-esterified homogalacturonans in the exinic outer layer and stigma available to function in fusion between the pollen and stigmatic ECM. To this end, we note that the contact point between the pollen and stigmatic ECM is characterized by a high density of epitopes recognized by JIM5 in Amborella and Illicium. In addition to the presumed function in pollen adhesion, we also emphasize the observation that the spatial array of homogalacturonans and AGPs within the microenvironment of the pollen grain adhesion zone provides a hydrophilic conduit for the movement of water from the stigma to the pollen grain and diffusion of molecules involved in cell-to-cell communication. The distribution of lipids and homogalacturonans within the adhesion zone mirrors the hydrophobic and hydrophilic boundary that is proposed to form a water gradient to which pollen grains and pollen tubes respond by hydrating and germinating directionally (Wolters-Arts et al., 1998, 2002).
Pollen tube growth along a stigmatic papilla also results in a zone of adhesion between the primary wall of these cells in Amborella and Illicium. A similar cell-to-cell attachment between the primary walls of the pollen tube and transmitting tissue has been reported for Arabidopsis (Lennon et al., 1998). As noted for the point of contact between the pollen grain and stigmatic ECM, this zone of adherence in Amborella and Illicium is characterized by a high concentration of low methyl-esterified homogalacturonans. The density of these molecules exceeds pre-pollination levels within the stigma ECM. We demonstrated that the cellular features of pollen tube tip growth and cell wall synthesis in Amborella and Illicium are similar to those reported for other angiosperms (Malhó, 2006). Therefore, the low methyl-esterified homogalacturonans at this zone of fusion between the primary wall of the pollen tube and transmitting tissue arise from de-esterification of the high methyl-esterified homogalacturonans secreted from the vesicle-rich pollen tube tip of both species. De-esterification of high methyl-esterified homogalacturonans may also be occurring within the transmitting tissue ECM. As hypothesized for the adhesion zone between the pollen grain and stigma, low methyl-esterified homogalacturonans are most likely functioning in cell-to-cell adhesion between the primary walls of the pollen tube and stigma within Amborella and Illicium.
Pollination and pollen tube growth result in significant remodeling of the stigmatic ECM structure in Amborella and Illicium. The cuticular layer of Amborella has a stark decrease in electron density and partial to complete loss of the reticulate cuticular network at the site of pollen adhesion, germination, and pollen tube growth within the fused cuticular ECM of adjacent stigmatic papillae. Similarly, the post-pollination transmitting tissue ECM in Illicium displays a localized decrease in the electron density and fibrillar nature of the primary wall that renders the primary wall frequently indistinguishable from the cuticular layer. Notably, there is an absence of the pre-pollination stigmatic cuticle where pollen tubes of both species are adherent to the primary wall of a stigmatic cell indicating the activity of cutinase. Remodeling of the stigmatic ECM with cutinase and other enzymes functions to support penetrative siphonogamous pollen tube growth within the transmitting tissue ECM and assists cellular contact between the two cell types to facilitate adhesion (guidance) and cell-to-cell communication (Hiscock and Allen, 2008).
In comparison to Amborella and Illicium in which the initial interactions between the pollen grain and the stigma ECM are similar to those of other species with a dry stigma following a compatible pollination, the interactions between the pollen grains and stigma ECM of Acorus are similar to those of other species with a dry stigma following a self-incompatible pollination. Pollen grains fail to hydrate as observed in the basal-relictual taxon Saururaceae (Pontieri and Sage, 1999) and species from monocots and eudicots with stigmatic SI (Sage et al., 2001; Hiscock and Allen, 2008). These unexpected results indicate that stigmatic SI may be present within the single population of Acorus that was used to confirm the pollen tube pathway following cross- and self-pollinations. The very limited number of successful cross-pollinations within the populations indicates that if SI is present, there are very few S-alleles potentially due to the clonal growth habit of Acorus (Vojtí
ková et al., 2004). Given the importance of SI in the evolution of breeding systems within angiosperms (de Nettancourt, 1997, 2001), rigorous studies will be required to provide definitive evidence of the presence or absence of SI in this or any other species of Acorus. Although stigmatic SI in association with bicellular pollen has been assumed to be rare, it is increasingly apparent that the phenomenon is more common than previously proposed (Sage et al., 2000). Stigmatic SI has traditionally been viewed to have evolved late in the history of angiosperms (Dickinson, 1995, Weller et al., 1995).
Is the transmitting tissue ECM of angiosperms homologous with the pollination droplet of extant gymnosperms?
The hypothesis that transmitting tissue ECM of angiosperms evolved from a pollination droplet homologous to that found in extant gymnosperms is based on (1) the proposed compositional and functional similarity of the pollination droplet to the freely flowing transmitting tissue ECM ("wet stigma") of some angiosperms and (2) the view that the wet stigma is ancestral to the cuticle-bound transmitting tissue ECM ("dry stigma") found in many angiosperms. We evaluated this hypothesis by phylogenetically tracing the history of these character states onto a recent angiosperm phylogeny (Stevens, 2001). Our analysis rejects the hypotheses as it resolves the dry stigma unequivocally as the ancestral state for extant angiosperms. Multiple, homoplasious occurrences of wet stigmas, in diverse groups of angiosperms, are reconstructed as having evolved from ancestors with dry stigmas. Our results also reject the related hypothesis that a gymnosperm-like pollination droplet produced by the carpel of a common ancestor of extant angiosperms conferred a selective advantage by using nectar-like stigmatic exudates to lure pollinating insects (Lloyd and Wells, 1992; Frohlich and Parker, 2000; Frame 2003a, b; Labandeira et al., 2007; Frohlich and Chase, 2007). Indeed, mapping the distribution of nectar rewards onto the angiosperm phylogeny does not resolve nectar production as ancestral for angiosperms (P. H. Weston, personal observation). As well, developmental factors promoting nectary development within the eudicots are not associated with either the stigma or nectariferous structures in extant basal-relictual angiosperms (Fourquin et al., 2005; Lee et al., 2005). The most recent common ancestor of extant angiosperms had a dry stigma and was pollinated either by wind or by pollen-collecting insects or both (Thien et al., 2003).
The presence of a prominent cuticle-bound exudate within the ovule micropyle of several basal-relictual angiosperm species also indicates that the ovule of the most recent common ancestor of extant angiosperms is unlikely to have had a freely flowing pollination droplet. It has been suggested that there is a close relationship between angiosperms and the extinct gymnosperm group Bennettitales (Crane, 1985; Doyle, 2006). Examination of the fossil record indicates that pollen grain capture and pollen tube growth likely occurred at the distal-most regions of a micropylar tube in Williamsonia (Bennettitales; Stockey and Rothwell, 2003). Pollen grains were not drawn into a pollen chamber where they subsequently germinated as occurs in many extant gymnosperms that form a pollination drop. Rather, pollen tubes in Williamsonia presumably grew from the distalmost region of the micropylar canal towards a nucellar plug (Stockey and Rothwell, 2003). Notably, the epidermis of the micropylar canal and nucellus had intact cuticles (Crane, 1985), indicating the absence of a freely flowing ECM. Finally, the micropyle of the Callospermarion-type ovule, a seed fern, is filled with a resinous exudate that may be bound by a cuticular boundary at pollen reception (Rothwell, 1977), suggesting that other early seed plants may have also lacked a freely flowing micropylar ECM.
What does the architecture of the transmitting tissue ECM in basal-relictual angiosperms indicate about the origin of transmitting tissue?
The cuticle functions as a permeability barrier against water loss from the epidermis and cuticular lipids play an essential role in this function (Jeffree, 2006). We propose that transmitting tissue evolved in concert with an increase in cuticle permeability that resulted from modifications in the biosynthesis and secretion of fatty acids needed for construction of the cuticle of an early carpel-like organ. Increased cuticle permeability would have enabled rapid, contact-driven male gametophyte development by exposing the pollen grain and pollen tube ECM to pre-existing molecules involved in pollen grain adhesion, hydration, germination, and pollen tube growth and guidance (methyl-esterified homogalacturonans, AGPs, lipids, LTPs, numerous esterases, LTPs). An acceleration of the reproductive cycle is cited as a critical innovation promoting the spectacular success of angiosperms (Stebbins, 1976; Takhtajan, 1976; Williams, 2008). Increased cuticle permeability would have also facilitated interactions between ECM molecules of the pollen and transmitting tissue ECM that operate in promoting species-specific recognition and compatibility. The ability of transmitting tissue to function in this context has also played a key role in outcrossing and thus, angiosperm success (de Nettancourt, 1997, 2001; Holsinger, 2000; Hiscock and Allen, 2008).
Dramatic modifications in cuticle structure and function are well known to occur as a result of mutations in genes needed for the biosynthesis and secretion of fatty acid molecules used in the construction of the cuticle in Arabidopsis and Zea (Lolle et al., 1992, 1997; Becraft et al., 1996; Sinha, 1998; Sinha and Lynch, 1998; Yephremov et al., 1999; Jin et al., 2000; Pruitt et al., 2000; Sieber et al., 2000; Chen et al., 2003; Broun et al., 2004; Kurdyukov et al., 2006; Li et al., 2007; Luo et al., 2007). Collectively, the cuticle phenotypes of mutant plants bear striking resemblance to features of the dry stigma of numerous angiosperms representing the lineages that diverged near the base of the phylogeny. In comparison to wild-type epidermal cells, which have a uniformly thickened and highly sculptured cuticle, mutant plants have a smooth cuticle proper and cuticular layer that is expanded and variable in thickness. Cuticular bosses, which develop in mutant plants (Sinha, 1998; Sieber et al., 2000), are rich in low methyl-esterified homogalacturonans (Sieber et al., 2000). Cuticle permeability in the mutant plants is enhanced and epidermal cells with the modified cuticle become postgenitally fused upon contact. Coherence of plant parts and enhanced cuticle porosity arise, in part, from pleiotropic effects of the mutations on the array of low methyl-esterified homogalacturonans within the cuticle (Sinha and Lynch, 1998; Sieber et al., 2000). The cuticle proper of mutant plants may be discontinuous or absent at the point of fusion wherein low methyl-esterified homogalacturonans function in cell adhesion. Most relevant to the present discussion is the observation that alterations in cuticle permeability of these mutant plants result in rapid, species-specific pollen grain adhesion, hydration, and germination as well as pollen tube growth on the epidermis of vegetative tissues (Lolle and Cheung, 1993; Sieber et al., 2000; Kurdyukov et al., 2006). Ectopic male gametophyte development on the vegetative epidermis is independent of a carpel-specific developmental program (Lolle et al., 1997).
We also hypothesize that modifications in the biosynthesis and secretion of fatty acid molecules used for construction of the cuticle of the carpel precursor enabled the production of the morphology commonly associated with the stigmatic surface (Heslop-Harrison and Heslop-Harrison, 1982). Cell division within the epidermis, which is typically anticlinal (Satina and Blakeslee, 1941), may become periclinal resulting in multicellular epidermal outgrowths in the aforementioned mutant plants (Becraft et al., 1996; Sieber et al., 2000). As well, epidermal cells in mutant plants can become enlarged and papillate (Becraft et al., 1996; Sinha, 1998; Jin et al., 2000; Sieber et al., 2000). Unicellular papillate stigmatic cells occur in some Nymphaeaceae, Illiciaceae, Austrobaileyaceae (Endress and Igersheim, 2000a), and Acoraceae (Buzgo and Endress, 2000). Multicellular stigmatic papillae that develop from periclinal divisions of epidermal cells are present in Amborella (Endress and Igersheim, 2000), Hydatellaceae (Rudall et al., 2007), Cabombaceae (Igersheim and Endress, 1997), and in some Nymphaeaceae (Igersheim and Endress, 1997) and Chloranthaceae (Endress, 1987; T. L. Sage, personal observations). Trimeniaceae develop multicellular stigmatic protrusions from periclinal and anticlinal divisions of epidermal cells (present study; Endress and Sampson, 1983). Although the transmitting tissue below the stigma epidermis has been interpreted as ground tissue in Trimenia (Bernhardt et al., 2003), it is possible that this transmitting tissue arises from periclinal and anticlinal divisions of cells originating in the carpel epidermis. The various stigmatic morphologies found in angiosperms function to enhance the epidermal surface area for pollen capture.
Conclusions
Structural and phylogenetic analyses from the present study have substantially revised traditional ideas regarding transmitting tissue origins in angiosperms. In contrast to a hypothesis that proposes that the transmitting tissue ECM of early angiosperms was homologous to the pollination droplet of gymnosperms, this study provides clear evidence in support of a hypothesis that the transmitting tissue ECM of the most recent common angiosperm ancestor was comparable in structure and function to a dry-type stigma. We demonstrate that this dry-type ECM was likely composed of a cuticle permeated by molecules deemed universally important for pollen grain adhesion, hydration, germination, and pollen tube growth. These molecules may have also been present in the pollen ECM and included homogalacturonans, AGPs, and lipids. Recent reviews on floral evolution emphasize how a potentially genetically simple trait, epidermal fusion, underpins floral developmental programs in angiosperms that result in the production of the tremendous diversity of floral and carpel morphology that is critical for promoting angiosperm success (Raven and Weyers, 2001). Here, we propose that processes leading to epidermal fusion, alterations in the biosynthesis and secretion of fatty acid molecules needed for construction of the cuticle of epidermal cells, were essential for promoting early angiosperm success because they resulted in the origin of a tissue that enables accelerated, species-specific compatible male gametophyte adhesion, hydration, and growth.
Numerous genes that have an impact on transmitting tissue differentiation and fusion within the carpel have been identified (Sessions and Zambryski, 1995; Roe et al., 1997; Bowman and Smyth, 1999; Heisler et al., 2001; Alvarez and Smyth, 2002; Kuusk et al., 2002; Azhakanadanam et al., 2008). One of these genes, CRABS CLAW (CRC) has recently been identified in Amborella (Fourquin et al., 2005). The expression of CRC is highest in the unfused dorsal epidermis of gynoecia in both Arabidopsis (Bowman and Smyth, 1999) and Amborella (Fourquin et al., 2005) and is downregulated during development in the sites of carpel fusion in Arabidopsis and the stigma of Amborella. Future studies assessing whether or not CRC and other genes played an essential role in transmitting tissue origins by modifying cuticle synthesis may provide important clues to the origin of transmitting tissue—a major floral innovation.
FOOTNOTES
1 This manuscript is dedicated to E. Heij, mentor. The authors thank G. W. Rothwell for helpful discussion, K. Sault for technical assistance, and A. Sage for editorial assistance. This research was funded by a Connaught New Faculty Award, University of Toronto start-up funds, and a Natural Sciences and Engineering Research Council of Canada grant to T.L.S. and a National Geographic Grant (#6974-01) to P.B. and T.L.S. 
4 Author for correspondence (e-mail: tammy.sage{at}utoronto.ca)
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0.05. Error bars represent 95% confidence interval.
