Pollen-pistil interactions and developmental regulation of pollen tube growth in Arabidopsis

In Arabidopsis, dehiscence of the anthers is usually accompanied by the transfer of pollen to the receptive surface of the pistil and the initiation of the process of self-pollination. Following pollen capture, a series of events are set in motion which culminate in the delivery of sperm cells to the female gametophyte and double fertilization. The critical events in this postpollination process relate to the establishment of a polarized growth pattern within the pollen grain, the elaboration of a pollen tube that extends by tip growth, the invasion of the tube into tissues of the pistil, and its directed growth towards the ovary as it homes in on its ovule targets. A major unresolved question in plant reproductive biology concerns the molecular basis of pollen tube guidance: how do pollen tubes navigate through tissues of the pistil and what signals guide them to the ovules?

The tissues of the pistil are uniquely specialized to promote these postpollination events (Hill and Lord, 1987; Gasser and Robinson-Beers, 1993). At the distal end of the Arabidopsis pistil is the stigma, a globular structure covered with a dense layer of papillar cells, the specialized epidermal cells that serve as receptacles for pollen grains. Below the papillar cells, the subepidermal cells of the stigma converge into a short solid style made up of a central core of transmitting tissue surrounded by a parenchymatous cortex and the epidermis. The style connects the stigma to the ovary, which consists of two locules containing ovules with haploid egg cells. In members of the Brassicaceae, including Arabidopsis and Brassica, contact with a papillar cell is a prerequisite for the activation of pollen and subsequent pollen tube development: pollen grains do not normally germinate on non-stigmatic surfaces (Lolle and Cheung, 1993). The cells of the transmitting tissue that line the path of pollen tubes are metabolically active secretory cells thought to provide the mechanical and nutritive environments for promoting normal pollen tube growth. Further, the high degree of precision in the directionality of pollen tube growth suggests that signals are continually being exchanged between pollen tubes and cells of the pistil that line their path. For example, a well established function of the pistil is in pollen recognition. In Brassicaceae, this recognition function is performed by the stigmatic papillar cells, as underscored by the observation that intra-and inter-specific incompatibility barriers operate at the level of the interaction between these cells and pollen in this family (Hiscock and Dickinson, 1993).

Recently, genetic ablation methodologies (Thorsness et al., 1993) and classical genetic approaches (Preuss et al., 1993) have been used to investigate the requirements for successful pollination in Arabidopsis. However, progress towards the molecular and/or biochemical identification of molecules that are essential for pollination as well as the genetic dissection of pollination processes requires a detailed understanding of post-pollination events as they occur in mature pistils and as a function of pistil and pollen development. Although the cyto-logical events that occur at the surface of the stigma following pollen capture have been described for Brassicaceae (Kanno and Hinata, 1969; Dickinson and Lewis, 1973; Elleman and Dickinson, 1990; Elleman et al., 1992), neither a detailed time-course of postpollination events nor a systematic description of the pollination responses of developing pistils and microspores are available for Arabidopsis. In this paper, we use light and electron microscopy to describe the responses of developing pistils and microspores of Arabidopsis thaliana in intra-and inter-specific pollinations.

The Arabidopsis thaliana strains used in the present study were RLD and C24. In Arabidopsis, the development of reproductive structures, pistils and pollen, occurs as a progression of floral buds and flowers along an inflorescence, with the youngest buds at the tip of the inflorescence and the more mature stages at its base. The ages of floral buds and flowers are conveniently expressed as follows: mature flowers at anthesis (flower opening) are designated as stage ‘0’, and younger buds containing progressively less mature structures are des ignated by consecutive numbers, with the bud immediately preceding the flower designated as ‘bud 1’, the next bud as ‘bud 2’, and so on.

Developing flower buds were dissected under a dissection microscope, pollinated with fresh pollen, and incubated on agar plates at 25°C. After different time intervals, the pollinated flower buds were fixed in ethanol:acetic acid (3:1) for approximately 30 minutes and softened in 1 N NaOH overnight. The samples were washed in distilled water, stained with decolorized aniline blue, squashed and examined with a Zeiss microscope equipped for UV epifluorescence. Each pollination was repeated at least three times.

Flower buds at different stages of development, corresponding to the stages used for analyzing pollen tube growth on developing pistils, were fixed under vacuum in 3% glutaraldehyde in 0.05 M cacodylate buffer (pH 7.2) for 3 hours at 20°C and then at 4°C for 3 hours. After rinsing in buffer, the samples were postfixed overnight in buffered 1% (w/v) OsO₄ at 4°C, washed in buffer, dehydrated on ice in a graded series of ethanol, and critical-point dried using liquid CO₂. The buds were then mounted on stubs and sepals and petals were removed carefully to expose the pistil and anthers. After coating with gold, the samples were examined in an AMR 1000 scanning electron microscope.

Samples were prepared for TEM by conventional chemical fixation. Pollinated tissues were fixed in 2.5% (v/v) glutaraldehyde in 0.05 M phosphate buffer (pH 7.15) at 20°C for 3 hours. They were then rinsed in buffer and further fixed in 1% (w/v) OsO₄ at 4°C overnight. After rinsing again in buffer, the samples were dehydrated through an acetone series and embedded in Spurr’s epoxy resin. Thin sections were cut with a diamond knife, stained with uranyl acetate and lead citrate, and observed in a Philips EM 300 electron microscope. Three replicates of each sample were processed. The conventional fixation method used here preserved the Arabidopsis pollen coat and was judged equivalent to the osmium vapour technique described by Elleman and Dickinson (1986).

In Arabidopsis, the entire process of postpollination events, starting from pollen deposition on the surface of the stigma and culminating with the entry of the pollen tube into the embryo sac, is rapid and is completed in 8-10 hours at 25°C. The path followed by pollen tubes within the Arabidopsis pistil is shown in Fig. 1. Examination of whole-mount preparations of pollinated flowers by fluorescence microscopy reveals that almost all of the pollen grains that come in contact with the papillar cells of the stigma surface germinate and produce pollen tubes that invade the stigma. Pollen tubes subsequently grow into the pistil and converge into the transmitting tissue of the style where they form a dense bundle (Fig. 1A). In the ovary, the pollen tubes fan out from the transmitting tissue and grow towards the ovules (Fig. 1A,B). To reach the embryo sac and effect fertilization, the pollen tube grows over the surface of the funiculus, a short stalk that anchors the ovule to the placenta, and enters through the micropyle (Fig. 1C). Interestingly, only one pollen tube is normally seen growing toward an ovule (Fig. 1C).

We examined the structure of the pollen-stigma interface by transmission electron microscopy at 5-minute intervals following hand-pollination of stigmas from mature flower buds. The pollen grain surface consists of a sculptured exine layer, in the cavities of which is embedded the tryphine – an extracellular coat rich in lipids, carbohydrates, and proteins derived from the sporophytic cells of the tapetum that lines the anther locules in which microspores develop. Fig. 2 shows the establishment, at the pollen-papillar cell interface, of an adhesion zone that appears to consist of the same material as the pollen tryphine. This adhesion zone is visible within 5 minutes after pollination (Fig. 3A), and persists through the onset of germination and the initiation of polar pollen tube growth (Fig. 3B-D). Within 10 minutes after capture, the pollen grain, which initially contains a relatively uniform distribution of organelles (Fig. 3A), undergoes a number of cytological changes that establish polarity in preparation for tube emergence; most obvious is the accumulation of vesicles at the germ pore near the site of adhesion (Fig. 3B,C). Within 20 minutes after capture, the emerging pollen tube with its vesicle-rich apical growth zone, can be seen invading the expanded papillar cell wall (Fig. 3D). The pollen tube subsequently continues its rapid growth down the length of the papilla between the separated outer and inner layers of the papillar cell wall (Fig. 4A,B), and by 45-50 minutes after pollination, pollen tubes have exited the papillar cell wall and have reached the intercellular matrix separating the papillar cells from the subepidermal cells of the stigma (Fig. 4C). This stage marks a transition in pollen-tube growth from elongation within the papillar cell wall to intercellular growth in the subepidermal zone of the stigma and in the transmitting tissue of the style and ovary.

Pollen grains are shed from the dehisced anthers of flowers in a ‘competent’ state and are capable of immediate activation and germination as soon as they come in contact with the receptive papillar cells of the stigma surface. To determine if microspores from the undehisced anthers of younger buds can interact productively with the stigmatic surface, we pollinated mature stigmas with microspores obtained from a sequence of immature floral buds, with the most mature bud designated as bud 1 (see Materials and Methods).

Developing microspores were obtained by dissection of undehisced anthers and were applied to pollen-free mature stigmas. Fig. 5 shows intact and dissected buds 1 and 2 (Fig. 5A,B), undehisced anthers removed from these buds (Fig. 5C), and the pollination behavior of microspores from bud 1 (Fig. 5D) and bud 2 (Fig. 5E). We found that hand-dissected bud 1 microspores were equivalent to mature pollen from dehisced anthers in their ability to adhere to papillar cells, germinate, and extend pollen tubes that invade pistil tissues (Fig. 5D). In contrast, pollen grains from bud 2 and younger buds neither adhered in large numbers nor germinated at the stigma surface (Fig. 5D). Thus, microspores acquire pollination competence during the period separating bud stages 1 and 2, which corresponds to approximately 6-8 hours in actual developmental time.

To determine if the Arabidopsis stigma is receptive to pollen throughout its development, we analyzed pistils from floral buds at progressively earlier stages of development ranging from anthesis to 5 days prior to anthesis (see Materials and Methods). A sequence of 13 consecutive buds corresponding to 13 developmental stages were assayed. Stigmas, or the distal end of undifferentiated pistils from immature buds, were manually exposed and pollinated with mature pollen grains. Pollinated pistils were incubated for 1 hour prior to microscopic examination to allow for pollen germination and pollen tube growth into the stigmatic tissue. Fig. 6 shows a dramatic developmental regulation of the ability of the pistil to support pollen development. Three developmental zones – mature, intermediate, and early – may be recognized along the inflorescence as inducing high, intermediate, and low pollen tube growth responses. In the mature zone consisting of buds 1, 2 and 3, the stigmas were highly receptive to pollen as evidenced by the adhesion and germination of a large number of pollen grains (198±25 (mean ± s.d.) grains/stigma), and by the rapid elongation of pollen tubes which, by 1 hour after pollination, had travelled approximately 200-250 µm to the base of the stigma (Fig. 6A-C). The stigmas of younger buds in the intermediate and early zones were progressively less efficient at supporting pollen tube growth. Fewer pollen grains adhered to and germinated on these immature stigmas: 83±28 (mean ± s.d.) grains/stigma were observed in buds from the intermediate zone and 24±15 (mean ± s.d.) grains/stigma in buds from the early zone. Further, the pollen tubes that were produced on these immature stigmas exhibited a much reduced growth rate: 1 hour after pollination, the tubes had extended only approximately 100 µm into the stigmas of buds 4 to 8 in the intermediate zone (Fig. 6D-H), and less than 50 µm into the stigmas of buds 9 to 13 in the early zone (Fig. 6I-M). Actual measurements of papillar cells and their initials show that they range in length from 60-90 µm in the mature zone, from 30-50 µm in the intermediate zone, and less than 30 µm in the early zone.

To determine if the inefficient growth of pollen tubes on immature stigmas was due to the inability of the tubes to exit the papillar cell wall or to a reduced growth rate, the pollination assays were carried out and pollen tube growth was allowed to proceed for a period of 12 hours before microscopic exami nation. We found that the tubes grew into the style and ovary after these prolonged incubations. The resulting patterns of pollen tube growth are shown for four representative bud stages in the fluorescence micrographs of Fig. 7 and are related to the morphology of the developing pistil in the scanning electron micrographs of Fig. 8.

Pollen tubes were found to follow their normal path through the transmitting tissue of the stigma, style, and ovary only in buds of the mature zone (Fig. 7A) in which pistils exhibited a defined stigma with elongated papillar cells (Fig. 8A) and contained mature ovules (see Fig. 1). In contrast, aberrant pollen tube growth was observed in immature pistils that exhibited incompletely differentiated stigmas and immature ovules. In buds of the intermediate zone in which a stigmatic zone with differentiating papillar cells caps the incompletely fused pistil (Fig. 8B), the majority of pollen tubes were still directed into the transmitting tissue, but a proportion of pollen tubes were observed outside this tissue, namely in the cortex of the ovary (Fig. 7B). In yet more immature buds of the early zone in which the cylindrical pistil exhibits, at its distal tip, only the epidermal initials of papillar cells (Fig. 8C) or a smooth undifferentiated epidermis (Fig. 8D), pollen tubes exhibited ectopic growth and were observed growing through the epidermis and parenchymatous cortex of the ovary as well as through the central region of the pistil (Fig. 7C,D). Interestingly, at this early stage of development, all epidermal surfaces of the pistil appeared to be receptive to pollen and allowed pollen tube entry (see Fig. 10A).

The direction of pollen tube elongation in the pistil was generally basipetal at all stages of bud development, and in buds from the early developmental zone, pollen tube elongation proceeded at the same rate whether the pollen tubes grew in the undifferentiated central region of the pistil or in the surrounding cortical tissue (Fig. 7C,D). The major difference between the patterns of pollen tube growth in early buds and mature buds was that pollen tubes were not directed towards the rudimentary ovules in immature pistils but rather continued to grow towards the base of the ovary (Fig. 7C,D).

The observations described above revealed a close correlation between the differentiation state of the pistil and the path followed by pollen tubes, and suggested that at maturity, cells of the pistil produce signals that are responsible for directing normal pollen tube growth. Under this hypothesis, after invading the pistil, the pollen tube would track first a positive signal emanating from cells of the transmitting tissue and then another signal emanating from the ovules. To determine if dif fusible signals are produced by the transmitting tissue, stigmas were excised with a sharp blade and pollen was applied directly onto the cut surface of the style (Fig. 9A). Microscopic examination of the pollinated styles after a 4-hour incubation showed that pollen grains had germinated and produced pollen tubes that grew randomly over the cut surface (Fig. 9B). However, basipetally directed growth was rarely observed and pollination of cut styles did not result in significant seed set: only one seed was obtained in ten independent pollinations.

As a rule, pollen grains will only adhere to, and germinate at, the surface of stigmatic cells in mature pistils. However, in pol linations of immature pistils, we occasionally observed pollen grains germinating on the epidermis of the ovary (see for example Figs 6L, 7D). To determine if other floral structures were also receptive to pollen early in floral bud development, we applied mature pollen grain to the surfaces of various floral organs from developing buds. Fig. 10 shows that in the early developmental zone (bud 9 and above), pollen grains germinated and produced pollen tubes on pistils, anthers, sepals, and petals. Interestingly, the epidermal surfaces of these organs were also receptive to pollen from another crucifer, Brassica oleracea (not shown). Further, the pollen tubes that were produced on these normally inhospitable surfaces were shown by ultrastructural analysis to invade the epidermal cell layer. Fig. 11A shows a pollen tube penetrating the ovary through the wall separating two epidermal cells with the site of contact apparently covered by pollen coat material. This ability of the epidermis of floral organs to support pollen tube development was observed mainly in the early developmental zone encompassing buds 9-13 above the first flower. In the intermediate zone, the floral epidermis allowed adhesion but not hydration and germination of pollen grains. Fig. 11B shows that pollen grains adhered to these surfaces via an adhesive zone probably consisting of pollen coat material in a manner similar to their legitimate adhesion to the papillar cell surface.

To analyze pollination responses in inter-specific crosses, we performed reciprocal crosses between Arabidopsis and Brassica. Pollen-pistil interactions in developing Brassica flowers are similar to those described here for Arabidopsis in that three developmental zones could be distinguished, based on competence to support pollen tube growth (unpublished observations). For each species, inter-specific pollinations were assayed in buds from these three developmental zones, and the outcome of these pollinations is shown in Fig. 12.

The behaviour of inter-specific pollen differed dramatically in mature pistils of the two species. On the mature Arabidopsis pistil, Brassica pollen germinated, invaded the papillar cells, and produced pollen tubes that grew into the underlying tissues of the pistil (Fig. 12A), albeit at lower efficiency than in intra-specific pollinations. Most significantly, Brassica pollen tubes did not grow towards Arabidopsis ovules. By contrast with the receptivity of Arabidopsis pistils to interspecific pollen, the mature Brassica pistil did not support the germination and subsequent development of Arabidopsis pollen (Fig. 12D): these inter-specific pollinations failed whether we used pistils from self-incompatible Brassica strains or a self-compatible strain that carries a null mutation at the self-incompatibility (S) locus (Nasrallah et al., 1994).

Interestingly, the ability to discriminate against interspecific pollen was apparently lacking at younger stages of pistil development: inter-specific pollinations were indistinguishable from intra-specific pollinations on immature pistils in both Arabidopsis (Fig. 12B,C) and Brassica (Fig. 12E,F).

By analyzing pollen tube growth at short time intervals beginning at 5 minutes after pollination, we have shown that postpollination events are very rapid in Arabidopsis. Cytological changes were evident within 5 minutes after pollen capture. The establishment of polarity within the pollen grain and pollen tube emergence occurred within 15 minutes after pollination, and 5 minutes later, pollen tubes had digested their way into the papillar cell wall. Pollen tubes were found to race into tissues of the pistil at a rate averaging approximately 4 µm per minute at 25°C.

Pollen tube growth through the tissues of the pistil was quantitatively and qualitatively related to the developmental state of pistil and pollen. During microspore development, the period separating bud 1 from bud 2, covering 6-8 hours, is identified as being critical for the acquisition by pollen grains of the competence to adhere to the stigma surface, germinate, and elaborate pollen tubes. However the nature of the maturation factors that are produced during this critical period is not known. Ultrastructural observations of the type presented in this paper and elsewhere (e.g. Elleman and Dickinson, 1990; Elleman et al., 1992) have pointed to the importance of the pollen coat in the pollen-papillar cell interactions that immediately follow pollen capture, particularly adhesion and hydration. The pollen coat is laid down over gametophytically encoded pollen wall components and consists in large part of sporophytically encoded molecules derived from the tapetum, a cell layer of the anther that acts as a nurse tissue for the developing microspores (Heslop-Harrison et al., 1973, 1974; Heslop-Harrison, 1975). One class of tapetum-derived molecules that are essential for pollen function in Arabidopsis was recently identified in genetic studies as long-chain lipids, which presumably act by stabilizing other critical components of the pollen coat (Preuss et al., 1993). The tapetum degenerates soon after the first mitotic microspore division at approximately the bud 7 or bud 8 stage in Arabidopsis (Toriyama et al. 1991; Thorsness et al. 1993), and the sporophytic domain of the pollen wall is, in all likelihood, already in place by the bud 2 stage. The bud 2-to-bud 1 developmental period is therefore likely to involve either the activation of sporophytically derived components or the synthesis and appropriate localization of gametophytically derived molecules, such as hydrolytic enzymes, implicated in the emergence of the pollen tube, that are known to be concentrated around the pores of pollen grains (Heslop-Harrison, 1975, 1987). In fact, it has been shown that microspores of tobacco and wheat isolated at the late-uninucleate and early binucleate stage respectively do mature in vitro, thus demonstrating that late pollen maturation events occur independently of the anther wall (Vicente et al., 1991). Whatever the nature of these pollen maturation factors, the failure of bud 2 pollen to grow on a mature stigma indicates that they are not synthesized or are not made accessible to the pollen grain by stigmatic cells.

Microspore maturation within the anther is accompanied by a parallel maturation of the pistil. We found that by the bud 3 stage, the pistil had acquired the competence to support abundant and normal pollen tube growth, whereas the incompletely differentiated younger pistils were less conducive to pollen tube growth. Based on the extent and nature of pollen tube growth in pistils at different stages of development, four phases of pollen tube growth can be delineated (Table 1). The first phase occurs at the pollen-papillar cell interface and includes the rapid deployment of adhesive forces, germination, establishment of polarity and the pattern of tip growth, and the breaching of the cuticle that covers the papillar cell. The second phase is characterized by pollen tube growth within the papillar cell wall. The third phase occurs in the stigma, style and ovary and consists of entry into the transmitting tissue and basipetally oriented intercellular growth. The fourth phase occurs in the ovary and is characterized by exit from the transmitting tissue and targeting towards the ovules.

Each of the four phases of pollen tube growth was dependent on the degree of differentiation of specific structures of the pistil. Thus, a much reduced number of pollen grains adhered to the stigma surface, germinated, and invaded the pistil when papillar cells were either unrecognizable as morphologically distinct cells or were only visible as papillar cell initials averaging less than 35% of mature length. In contrast, pistils with differentiated papillar cells that had expanded to 75% of their final length induced efficient pollen tube development. The papillar cells of these mature-stage pistils presumably contain factors that promote pollen germination and tube growth. Although information on the nature of such stimulatory factors is lacking, papillar cell-specific proteins that potentially function in pollination have been identified in Arabidopsis (Dwyer et al., 1992; Dwyer et al., personal communication). In addition, recent experiments in petunia (Mo et al., 1992; Ylstra et al., 1994), tobacco (Ylstra et al., 1992), and maize (Franken et al., 1991) have demonstrated that flavonols are required at low concentration for pollen germination and can be supplied to the pollen grain by the pistil. With the availability of Arabidopsis mutants carrying defects in the phenylpropanoid pathway, it should be feasible to test whether flavonols or their derivatives are effective in stimulating pollen germination in this species as well.

Interestingly, pollen tubes elongated at a reduced rate, lacked directionality, and grew randomly within the cortical tissue of pistils in the early developmental stages, when the central core of cells that eventually form the transmitting tissue was still exposed in the incompletely formed pistil and when the ovules were still rudimentary structures. Several hypotheses have been proposed concerning the nature of the signals reponsible for the directed growth of pollen tubes within the pistil (reviewed by Mascarenhas, 1993). Our analysis indicates that, whatever their nature, these signals are subject to strict developmental regulation. Thus, the critical papillar cell factors are apparently synthesized and incorporated into the cell wall by the time these cells have completed approximately 30% of their expansion. This conclusion is supported by the results obtained with genetically ablated Arabidopsis pistils (Thorsness et al., 1993) in which pollen tubes elaborated by wild-type pollen were properly directed into the papillar cell wall and into the underlying tissues, even though the biochemically inactive papillar cells were expanded to only approximately 30% of wild-type papillar-cell length at maturity.

For their part, the signals that direct the pollen tube into the transmitting tissue and towards the ovules are either first emitted by the appropriate cells of the transmitting tract and ovules during the mature developmental stage or reach physiologically relevant concentrations at that time. The observation that only one pollen tube grows toward an ovule in the mature pistil implicates the action, not only of positive signals, but also of signals inhibitory to supernumerary pollen tubes. Whether inhibitory signals are also produced by cortical cells at pistil maturity to prevent pollen tube growth in the intercellular space of the cortex is not known. However, it is evident that these cortical cells lack the capacity to induce directional pollen tube growth. Pollen tubes that germinated on the surface of cut styles grew in a random fashion, underscoring the impor tance of stigmatic cells, perhaps papillar cells, in establishing pollen tube directionality.

An unexpected observation of our studies was that the epidermal surfaces of floral structures at early stages of development had the capacity to support pollen tube growth. Previously, pollen germination and pollen tube growth on nonstigmatic surfaces had only been described for the shoot epidermis of the fiddlehead (fdh) mutant (Lolle and Cheung, 1993). The FDH gene was suggested to encode a factor that normally restricts competence to support pollen tube growth to the stigmatic region of the pistil. The observations reported here indicate that receptivity to pollen tube growth is not restricted to the stigma in wild-type plants but is a general property of the floral epidermis at early stages of flower development. Presumably, the ability of epidermal cells to produce factors that promote pollen tube growth are part of a floralspecific program that is activated during floral morphogenesis and later restricted to stigmatic epidermal cells as papillar cells differentiate and complete their maturation. The fdh mutation would perturb this developmental restriction by maintaining the floral epidermis in an immature state and expanding this floral-specific program into the vegetative domain.

An important function assigned to the stigma in crucifers is the discrimination against pollen grains from other species. Hiscock and Dickinson (1993) described the unilateral incom patibility of reciprocal inter-specific crosses between Arabidopsis and a self-incompatible Brassica oleracea strain. We characterized inter-generic pollination responses further by examining pollen tube growth patterns in developing pistils and by performing reciprocal crosses between Arabidopsis and a range of Brassicas including self-compatible and self-incom-patible strains. In keeping with the results of Hiscock and Dickinson, we found that mature as well as immature Arabidopsis pistils supported the development of Brassica pollen tubes, whereas only immature pistils of Brassica supported the germination and tube development of Arabidopsis pollen. We further showed that in very young buds of either species, the growth pattern exhibited by inter-specific pollen tubes was indistinguishable from that of intra-specific tubes. Interestingly however, in the ovary of mature Arabidopsis pistils, the growth patterns of Brassica and Arabidopsis pollen tubes differed markedly, and Brassica tubes were not directed towards the ovules. Thus, inter-specific incompatibility apparently operates at the level of the ovary in Arabidopsis, with Brassica pollen tubes unable to perceive or respond to the signals emanating from the ovules. In contrast, inter-specific incompatibility operates at the level of the stigma in Brassica, with the papillar cells actively rejecting Arabidopsis pollen or simply lacking the relevant germination-promoting factors.

It should be noted that because of cytological and developmental similarities, pollen rejection in inter-specific unilateral incompatibility has been proposed to be due to the functioning of genes operative in intra-specific incompatibility (Hiscock and Dickinson, 1993). Under this proposal, Arabidopsis pollen would be recognized and rejected by the Brassica stigma through the action of genes encoded in the self-incompatibility (S) locus, and specifically as a consequence of signaling by the S-locus receptor kinase (SRK) and S-locus glycoprotein (SLG) genes, a gene pair expressed in Brassica papillar cells and required for the operation of self-incompatibility in this genus (Nasrallah and Nasrallah, 1993). However, our observation that a self-fertile Brassica strain that has a defective SRK gene allows the growth of self pollen tubes but is nevertheless still able to reject Arabiodpsis pollen does not support a role for S-locus genes in inter-specific incompatibility. Rather, these results suggest that at least in crucifers, interspecific and intra-specific pollen recognition phenomena are dependent on distinct signaling pathways.

This work was supported by grant No. IBN-9220401 from the National Science Foundation.