The structural events in the stigma and transmitting tissue of Petunia hybrida pistils that accompany compatible and incompatible intraspecific pollinations have been investigated in detail, together with the changes in reserve levels that also take place at this time. Many of these phenomena may be explained in terms of 3 phases of secretion by the cells in the upper regions of the transmitting tissue. The first, independent of pollination, results in the deposition of an intercellular matrix, rich in protein and carbohydrate. The second, triggered by pollination, although independent of the compatibility of the pollen grain, involves synthesis of molecules believed to be specific to the S(incompatibility)-gene system. The third phase of secretion occurs only following a compatible mating, and involves the transfer of stylar reserves to support the growth of the pollen tubes. These observations are discussed in terms of current models of the incompatibility mechanism operating in Petunia.

In plants with gametophytic control of pollen compatibility with respect to the style, much attention has been focussed upon the structure and physiology of the pistil. For example, the cytology of the stigma of Petunia sp. (Konar & Linskens, 1966a) and that of the stylar transmitting tissue in a variety of species (van der Pluijm & Linskens, 1966; Sassen, 1974; Bell & Hicks, 1976; Cresti, Went, Pacini & Willimse, 1976) has been the subject of recent intensive investigation. Likewise, the stigma and style of Lilium, a plant with a stylar canal rather than a transmitting tissue per se, have also been described in detail (Rosen & Thomas, 1970). Physiological studies, however, have extended into the differences in stylar metabolism following compatible and incompatible crosses. Bredemeijer (1974) has demonstrated differences in the stylar peroxidase isozymes following such crosses in Nicotiana, while different levels of RNA and protein synthesis have been described following crosses of different compatibility in Petunia by van der Donk (1974).

Structural aspects of these stylar differences have yet to be fully examined. Certainly, structural changes accompanying cross and self-pollination have been described in Oenothera (Dickinson & Lawson, 1975) but here few alterations appear to occur in the female tissue. In any event, this species possesses neither an organized transmitting tissue, nor a stylar canal, making localization of the site of the self-incompatibility reaction difficult. Likewise, although Lycopersicum has a compact style with a transmitting tissue, the many elegant ultrastructural investigations into the differences between compatible and incompatible crosses (e.g. de Nettancourt et al. 1973) have not considered stylar changes in detail. Our knowledge of the reaction of compact styles to pollination is thus restricted to studies of the disruption of the tissue by pollen tube growth in Lychnis (Crang, 1966), and measurement of uptake of materials from the style by pollen tubes in a number of species (Linskens & Esser, 1959; Kroh, Miki-Hirosige, Rosen & Loewus, 1970; Kroh & Helsper, 1974). Stylar changes may, however, be of considerable importance since little is known of the factors that induce relatively slow growth of the incompatible tubes, when compared with the rate of compatible tube extension. While structural data may be few, models explaining these events certainly are not. Kroes (1973), for example, suggests that incompatible tubes lack an enzyme necessary to use the nutrients available in the style while, in an elegant hypothesis for Petunia, van der Donk (1975) proposes that, following a pollination-generated stimulus, synthesis of polypeptides takes place in the style. It is the interaction between these ‘pollination-polypeptides’ (or a larger molecular assembly containing them) and the tube that results in the ‘compatible’ or ‘incompatible’ style of growth.

In an attempt to align some of these hypotheses with structural events, we describe here the changes in the stigma and style accompanying compatible and incompatible intraspecific pollination in Petunia.

Plant material

Seedlings homozygous with respect to the incompatibility genes were raised by means of bud pollination of clones Ka3 and Tzu (genotypes 22 and 33 respectively), the seeds of which were originally kindly supplied by Professor H. F. Linskens. Following growth and flowering in the greenhouse, individual flowers were detached for the pollination studies. These experiments were carried out in a well-lit air-conditioned chamber, maintained at a constant temperature of 25 °C.

Studies of pollen tube growth

Pollen tube growth was measured using material stained with aniline blue examined under a Leitz epi-UV irradiance system fitted to a Leitz Dialux microscope. Styles were prepared following a modifications of Linskens & Esser’s (1957) technique. Material was first softened for I min at 100 °C in a 5 % aqueous solution of sodium sulphite, and then stained in a 0·1 % solution of aniline blue ??O?N K3PO4, for 5 min. The tissues were examined mounted in the stain.

Determination of starch levels

With the aid of a sharp scalpel short sections were excised from different regions of the style and, using light pressure, the transmitting tissue contained in the section extruded onto a clean slide. Difficulties were encountered in extracting the very top of the transmitting tissue, that which interfaces with the stigma. However, by shaving off the stigmatic papillae with a microscalpel it was possible to expose, and subsequently express, the top layer of the transmitting tissue.

Starch in these cells was then stained with a saturated solution of iodine in potassium iodide for 5 min at room temperature. Following rinsing in 0·3 M Sorensen phosphate buffer, the transmitting tissue was mounted in glycerol, squashed carefully, and the preparation sealed with rubber solution. The intensity of the stain was measured using a Vickers M85 scanning microdensitometer, operating at a wavelength of 500 nm. The method of preparation produced a monolayer of stained cells, and readings were taken of a constant area of this monolayer, limited by the field of the densitometer ‘mask’. Each field contained between 4 and 6 cells. It must be emphasized that the readings resulting from this work can in no way be linked stoichiometrically with the starch in the tissue, but they do provide a useful indication of the levels of this reserve. In some of the results it was considered helpful to express the data as ‘amounts per flower’. To obtain this measure, the average was taken of the readings from each level in the style.

Preparation for light and electron microscopy

For enzyme cytochemistry, fresh material was sectioned at 5 μm in a Slee cryostat and stained to reveal the location of acid phosphatase (Knox & Heslop-Harrison, 1970), esterase (Pearse, 1960) and peroxidase (Jensen, 1962).

Tissue for electron microscopy was prepared according to Dickinson & Lawson (1975). To identify protein in this material, sections were digested with protease (Dickinson & Potter, 1975). Thick sections (1·5 μm) of these Epon blocks were also cut for light microscopy. Carbohydrate was localized in these sections using treatment with periodic acid and Schiff’s reagent (Feder & O’Brien, 1968).

The stigma

At anthesis, degeneration of the stigmatic tissue takes place completely independently of pollination and indeed, the compatibility of that pollination. Prior to this event, electron micrographs reveal the papillae not only to contain reserves of starch and saturated lipid, but also plastids (see Figs. 1, 2), dictyosomes, associated vesicles, microbodies and mitochondria. As degeneration of these commences, large droplets of lipid become evident inside the cytoplasm of a proportion of the cells (see Fig. 3). The plasma membranes of these cells then rupture, releasing the lipid to coalesce with other extracellular droplets (see Figs. 4, 5), and the remaining cytoplasmic content to form small islets of degenerate cytoplasm that float in the lipid (see Fig. 3). Cytochemical tests, however, show the stigmatic extract to contain sugars and also acid phosphatase. This enzyme is not distributed evenly, but apparently contained in droplets in the stigmatic fluid (see Fig. 6). No reaction was observed with the cytochemical tests for either peroxidase or esterase.

Fig. 1.

Young stigmatic papillar cells of Petunia hybrida containing a full complement of organelles, including mitochondria (m) and starch-containing plastids (p). Droplets of saturated lipid (l) are also present in this cytoplasm. Fig. 1, × 7540; Fig. 2, × 5256.

Fig. 1.

Young stigmatic papillar cells of Petunia hybrida containing a full complement of organelles, including mitochondria (m) and starch-containing plastids (p). Droplets of saturated lipid (l) are also present in this cytoplasm. Fig. 1, × 7540; Fig. 2, × 5256.

Fig. 2.

Young stigmatic papillar cells of Petunia hybrida containing a full complement of organelles, including mitochondria (m) and starch-containing plastids (p). Droplets of saturated lipid (l) are also present in this cytoplasm. Fig. 1, × 7540; Fig. 2, × 5256.

Fig. 2.

Young stigmatic papillar cells of Petunia hybrida containing a full complement of organelles, including mitochondria (m) and starch-containing plastids (p). Droplets of saturated lipid (l) are also present in this cytoplasm. Fig. 1, × 7540; Fig. 2, × 5256.

Fig. 3.

Degenerating stigmatic papillae showing accumulations of unsaturated lipid (l) in intact protoplasts (p). Other cells (c) have completely degenerated to form lipid droplets and vesicles (arrows) containing fibrogranular material. × 3078.

Fig. 3.

Degenerating stigmatic papillae showing accumulations of unsaturated lipid (l) in intact protoplasts (p). Other cells (c) have completely degenerated to form lipid droplets and vesicles (arrows) containing fibrogranular material. × 3078.

Fig. 4.

Scanning electron micrograph of stigmatic surface. Individual papillae (s) protrude through the stigmatic fluid. The spheres visible (arrows) may represent the droplets of unsaturated lipid visible in Fig. 5. × 770.

Fig. 4.

Scanning electron micrograph of stigmatic surface. Individual papillae (s) protrude through the stigmatic fluid. The spheres visible (arrows) may represent the droplets of unsaturated lipid visible in Fig. 5. × 770.

Fig. 5.

Transmission electron micrograph of material depicted in Fig. 4. A proportion of the papillae (s) remain intact and are invested by the lipidic residues of the degenerate cells, × 3348.

Fig. 5.

Transmission electron micrograph of material depicted in Fig. 4. A proportion of the papillae (s) remain intact and are invested by the lipidic residues of the degenerate cells, × 3348.

Fig. 6.

Light-microscopic preparation treated to reveal acid phosphatase. The enzyme is localized in small droplets which occur both on the papillar surfaces (s) and in the large spaces between the loosely packed cells of the stigma. × 1176.

Fig. 6.

Light-microscopic preparation treated to reveal acid phosphatase. The enzyme is localized in small droplets which occur both on the papillar surfaces (s) and in the large spaces between the loosely packed cells of the stigma. × 1176.

It is into this lipid-rich fluid that the pollen grain alights. Within 30 min it has germinated, and the pollen tube begun to grow between the papillae (see Figs. 7, 8) into the stigma.

Fig. 7.

Scanning electron micrograph of a pollen grain germinating on the stigma. The pollen tube (t) has penetrated the stigmatic surface (arrow). A papilla (s) is also visible, as is a tube (t1) from another grain. × 1534.

Fig. 7.

Scanning electron micrograph of a pollen grain germinating on the stigma. The pollen tube (t) has penetrated the stigmatic surface (arrow). A papilla (s) is also visible, as is a tube (t1) from another grain. × 1534.

Fig. 8.

Transmission electron micrograph of material shown in Fig. 7. The pollen tube (t) is growing down between stigmatic papillae (S) invested by droplets (arrows) of the lipidic remains of degenerated cells. × 6016.

Fig. 8.

Transmission electron micrograph of material shown in Fig. 7. The pollen tube (t) is growing down between stigmatic papillae (S) invested by droplets (arrows) of the lipidic remains of degenerated cells. × 6016.

The cells of the transmitting tissue

The cells of this region present the features of normal somatic meristematic plant cells, albeit with rather thick walls, some of which may measure up to 0·5 μm in width. Apart from the conventional cell contents, the cytoplasm of these cells possess reserves of lipid and starch, considerable numbers of dictyosomes, mitochondria, and a large endoplasmic reticulum (see Fig. 9). Frequently, these cells contain microbodies. Close examination of the transmitting tissue with the electron microscope reveals it to be differentiated into 2 distinct regions, each with a characteristic cell morphology. The tissue in the ‘neck’ of the style contains large spherical cells (see Fig. 10), possessing characteristic ridges in their walls (see Fig. 11) which cause them to ‘key’ into an adjacent cell. In the remainder of the transmitting tissue, the cells are elongate, more loosely packed and do not have superficial ridges. The organelle content of both types of cell is similar, but the ‘neck’ cells often appear to contain more endoplasmic reticulum and ribosomes. The endoplasmic reticulum, be it in the neck cells or elsewhere in the transmitting tissue, is regularly associated with vesicles (see Fig. 12). These vesicles are also frequently observed subjacent to, or apparently merging with the plasma membrane.

Fig. 9.

Transmitting tissue cell of Petunia showing a starch-containing plastid (p), mitochondria (m), a dictyosome (d) and elements of the endoplasmic reticulum (arrows). × 18165.

Fig. 9.

Transmitting tissue cell of Petunia showing a starch-containing plastid (p), mitochondria (m), a dictyosome (d) and elements of the endoplasmic reticulum (arrows). × 18165.

Fig. 10.

The ‘neck’ region of the transmitting tissue. The cells of this region are spherical and possess unusual ‘key’ junctions (arrows) with their neighbours (shown better in Fig. 11). Note also the electron-opaque intercellular spaces. × 1960.

Fig. 10.

The ‘neck’ region of the transmitting tissue. The cells of this region are spherical and possess unusual ‘key’ junctions (arrows) with their neighbours (shown better in Fig. 11). Note also the electron-opaque intercellular spaces. × 1960.

Fig. 11.

Protease-digested material similar to that shown in Fig. 10. The loss of electron opacity of the intercellular spaces is striking. The ‘key’ junctions between cells (arrows) are also conspicuous in this micrograph. × 4504.

Fig. 11.

Protease-digested material similar to that shown in Fig. 10. The loss of electron opacity of the intercellular spaces is striking. The ‘key’ junctions between cells (arrows) are also conspicuous in this micrograph. × 4504.

Fig. 12.

Portion of cell in ‘neck’ region of the transmitting tissue. Elements of the endoplasmic reticulum are evident (e) as are associated vesicles (arrow). × 25 500.

Fig. 12.

Portion of cell in ‘neck’ region of the transmitting tissue. Elements of the endoplasmic reticulum are evident (e) as are associated vesicles (arrow). × 25 500.

Particularly striking are the large intercellular spaces characteristic of the transmitting tissue, which appear to contain an electron-opaque matrix (see Fig. 10). Again differences occur between cells in the neck of the style and those in the remainder of the tissue, for the content of the spaces formed between the ‘neck’ cells is particularly sensitive to protease digestion (see Fig. 11), while that between cells elsewhere in the transmitting tissue remains unaffected, even following long periods of digestion. Cytochemical investigation of the intercellular spaces also reveals differences between the ‘neck’ and lower regions of the tissue, for, while acid phosphatase (see Fig. 13), peroxidase, and carbohydrate (see Fig. 14) are common to spaces throughout the tissue, the neck cells are surrounded by spaces containing esterase, in addition to generally larger amounts of these other constituents.

Fig. 13.

Light-microscopic preparation of ‘neck’ region of transmitting tissue, treated to reveal acid phosphatase. Note the presence of this enzyme in the intercellular spaces. × 1178.

Fig. 13.

Light-microscopic preparation of ‘neck’ region of transmitting tissue, treated to reveal acid phosphatase. Note the presence of this enzyme in the intercellular spaces. × 1178.

Fig. 14.

As Fig. 13, but material reacted to show the location of carbohydrate. Starch (arrows) is visible within the cells, but the intercellular matrix is also PAS-positive. × 761.

Fig. 14.

As Fig. 13, but material reacted to show the location of carbohydrate. Starch (arrows) is visible within the cells, but the intercellular matrix is also PAS-positive. × 761.

While all the preceding events take place in virgin styles, other changes do overcome the ‘neck’ cells of the transmitting tissue on pollination, independent of the compatibility of the mating. The first indications of this stylar reaction to pollination is a rise in the number of polyribosomes in these cells (see Fig. 15). This takes place between 0·5 and 2 h after pollination and is accompanied by large numbers of single-membraned cytoplasmic inclusions becoming associated with the plasma membrane to form a characteristic ‘embayment’ (see Fig. 16). These embayments may measure up to 0·2 μm in the maximum dimension and contain a grey fibrillar matrix, which merges with that of the wall. This activity at the plasma membrane is comparatively short-lived and, within about 4 h, this membrane has returned to its original aspect.

Fig. 15.

Cells of the ‘neck’ region of the transmitting tissue 2 h after pollination. Large numbers of polyribosomes (arrows) are present and the embayments (e) are also visible. × 27675.

Fig. 15.

Cells of the ‘neck’ region of the transmitting tissue 2 h after pollination. Large numbers of polyribosomes (arrows) are present and the embayments (e) are also visible. × 27675.

Fig. 16.

As Fig. 15. The embayments (e) of the plasma membrane are particularly striking, as is their fibrillar content. × 14287.

Fig. 16.

As Fig. 15. The embayments (e) of the plasma membrane are particularly striking, as is their fibrillar content. × 14287.

All these events occur before the arrival of the pollen tubes, which are at this time growing at a rate of about 150 μm/h through stigmatic tissue. Close to these extending pollen tubes many of the ‘neck’ cells of the transmitting tissue appear necrotic, containing dark granular cytoplasm, large numbers of vesicles, and disorganized plastids (see Fig. 17). In addition, cytoplasmic fragments may frequently be observed in the intercellular spaces dividing these cells (see Fig. 18). These necrotic cells, if tested some 24 h after pollination, react most strikingly with protease, appearing to be almost totally sensitive to the enzyme (see Fig. 19).

Fig. 17.

Pollen tube (t) growing through the ‘neck’ region of the transmitting tissue. Near the tube some of the cells (n) appear disorganized and necrotic, while others (c) display a normal aspect. × 5771.

Fig. 17.

Pollen tube (t) growing through the ‘neck’ region of the transmitting tissue. Near the tube some of the cells (n) appear disorganized and necrotic, while others (c) display a normal aspect. × 5771.

Fig. 18.

‘Neck’ region of transmitting tissue during passage of pollen tubes. Intact cells (c) are present, but also visible is a degenerate protoplast (n) and cytoplasmic fragments (f) × 5769.

Fig. 18.

‘Neck’ region of transmitting tissue during passage of pollen tubes. Intact cells (c) are present, but also visible is a degenerate protoplast (n) and cytoplasmic fragments (f) × 5769.

Fig. 19.

Degenerate protoplast (n) as shown in Fig. 18, following digestion with protease. This cell is almost totally sensitive to the enzyme, whereas intact cells (c) are not. × 7185.

Fig. 19.

Degenerate protoplast (n) as shown in Fig. 18, following digestion with protease. This cell is almost totally sensitive to the enzyme, whereas intact cells (c) are not. × 7185.

The stylar cells following self- and cross-pollinations

Although the cells of all regions of the transmitting tissue appear to react independently of the compatibility of the pollen tube approaching them, their behaviour following its passage differs considerably according to the nature of the mating.

Following a compatible cross, for example, cells are characterized by large vacuoles invested by a thin peripheral layer of cytoplasm. This cytoplasm is no longer rich in reserves, but instead contains a nucleus, few mitochondria and scattered dictyosomes (see Figs. 20, 21). The transmitting tissue cells of self-pollinated flowers, on the other hand, resemble those of unpollinated flowers. Little depletion of reserves is indicated by the micrographs (see Fig. 22), and the cytoplasm: vacuole ratio remains largely unchanged.

Fig. 20.

Tangential section of a transmitting tissue cell following the passage of compatible pollen tubes. Note the absence of reserves from this protoplast. × 9585.

Fig. 20.

Tangential section of a transmitting tissue cell following the passage of compatible pollen tubes. Note the absence of reserves from this protoplast. × 9585.

Fig. 21.

As Fig. 20, but median section revealing the large vacuoles (v) of these cells. × 6140.

Fig. 21.

As Fig. 20, but median section revealing the large vacuoles (v) of these cells. × 6140.

Fig. 22.

Transmitting tissue cell following the passage of incompatible pollen tubes. Note the presence of starch (st) in the plastids and droplets of lipid (l) free in the cytoplasm. × 9060.

Fig. 22.

Transmitting tissue cell following the passage of incompatible pollen tubes. Note the presence of starch (st) in the plastids and droplets of lipid (l) free in the cytoplasm. × 9060.

Starch metabolism during pollination

While the changes above may be easily observed qualitatively, a quantitative measure of the events is not so simply obtained. Nevertheless, microdensitometry of preparations in which the starch has been stained (see Fig. 23) provides a quantitative representation of levels of this reserve.

Fig. 23.

Light-microscopic preparation of lower region of the transmitting tissue treated with PAS to reveal the presence of starch. Although the cell walls react and the intercellular regions are slightly sensitive, the main staining is in the starch grains of the stylar cells. × 934.

Fig. 23.

Light-microscopic preparation of lower region of the transmitting tissue treated with PAS to reveal the presence of starch. Although the cell walls react and the intercellular regions are slightly sensitive, the main staining is in the starch grains of the stylar cells. × 934.

From such results, it is clear that in unpollinated flowers no change occurs in the starch content of the transmitting tissue until the final degeneration of the style. After a cross-pollination, however, a massive decrease in starch levels takes place, while, following a self-mating, there is only a slight depletion of the reserve (see Table 1).

Table 1.

Levels of IKI-stainable material, expressed as arbitrary microdensitometer units per flower. (The standard errors are shown in parentheses.)

Levels of IKI-stainable material, expressed as arbitrary microdensitometer units per flower. (The standard errors are shown in parentheses.)
Levels of IKI-stainable material, expressed as arbitrary microdensitometer units per flower. (The standard errors are shown in parentheses.)

In order to investigate a possible relationship between the passage of the pollen tubes and depletion of the starch, the number of pollen tubes at different levels in the styles was expressed as a percentage of the number of pollen grains present, and this percentage considered in terms of the starch levels in the entire pistil. When these results are plotted (see Fig. 24) a clear relationship may be detected in compatible crosses, the starch of the stylar cells decreasing with the passage of the pollen tubes through the adjacent intercellular spaces. Following incompatible crosses, this inverse relationship is not nearly as marked.

Fig. 24.

The number of pollen tubes and levels of starch at different points in the style 24 h after pollination following compatible (A), and incompatible (B) pollinations. The pollen tube number (•) is expressed as a percentage of the grains on the stigma surface. The levels of starch (▪) are in arbitrary microdensitometer units.

Fig. 24.

The number of pollen tubes and levels of starch at different points in the style 24 h after pollination following compatible (A), and incompatible (B) pollinations. The pollen tube number (•) is expressed as a percentage of the grains on the stigma surface. The levels of starch (▪) are in arbitrary microdensitometer units.

This difference in starch metabolism between tubes of differing compatibility is particularly well underlined by a series of readings taken at the neck of the transmitting tissue, subjacent to the stigma, when pollen tube distribution is almost identical (see Fig. 25 A, E). Starch levels in the selfed styles are far in excess of those found following cross-pollination.

Fig. 25.

Levels of IKI-stained starch, expressed in arbitrary microdensitometer units, in unpollinated (▫), selfed (▨), and crossed (▩) flowers at 4 levels in the pistil, 24 (A,B,C,D) and 48 (E,F,G,H) h after pollination. The pollen tube numbers are given in brackets under the lower axes. The regions of the pistil from which measurements were taken are as follows: ?, E, stigma; B, F, top quarter of style; c, G, second quarter; and D, H, third quarter.

Fig. 25.

Levels of IKI-stained starch, expressed in arbitrary microdensitometer units, in unpollinated (▫), selfed (▨), and crossed (▩) flowers at 4 levels in the pistil, 24 (A,B,C,D) and 48 (E,F,G,H) h after pollination. The pollen tube numbers are given in brackets under the lower axes. The regions of the pistil from which measurements were taken are as follows: ?, E, stigma; B, F, top quarter of style; c, G, second quarter; and D, H, third quarter.

In addition to these effects of pollination, measurement of starch content at 4 levels at 24 and 48 h after pollination indicates that in styles pollinated with grains of either compatability, a detectable ‘wave’ of starch synthesis precedes the tube tip in its passage through the transmitting tissue (see Fig. 25 C, D).

The maturing stigma and first events after pollination

The final stages in stigmatic maturation which appear to involve the ‘degeneration’ of the papillae (Dumas, 1975) are clearly independent of pollination and, indeed, the compatibility of pollination. Konar & Linskens (1966b) have proposed that the stigma acts purely as a location for the germination of pollen and is not involved in its nutrition per se, and there is little from the present investigation to indicate otherwise.

However, while the pollen grains are germinating in this stigmatic exudate, striking changes appear to be induced in the subjacent regions of the transmitting tissue. These start with a marked increase in polyribosome number in the cells, and continue with the formation of the characteristic ‘embayments’. While conclusive evidence is, of course, unavailable, these profiles do indicate a transfer of cytoplasmically synthesized material into the carbohydrate and pectin rich intercellular spaces (Kroh, 1973; Kroh & van Bakel, 1973; Sassen, 1974). The nature of the stimulus that travels to these cells of the stylar neck and triggers these changes is not clear. Linskens & Spanjers (1973) have recorded a difference in the electrical resistance of the pistil after pollination, but it remains equally possible that these changes are induced by a flow of chemical messenger. These events must also be accompanied by biochemical changes, such as the activation of the glutamic dehydrogenase described by Roggen (1967).

Biochemical studies (van der Donk, 1975) have indicated pollination to stimulate the production of ‘recognition’ polypeptides, specific to the incompatibility (S) genotype of the plant. It is striking that this aggregation of ribosomes into polysomes and the apparent secretion of materials by the cells in the neck of the transmitting tissue is so closely synchronized with the proposed synthesis of the s-specific polypeptides, all these events occurring in the style ahead of the advancing pollen tubes.

Once the pollen tubes reach the transmitting tissue, there is little doubt that some cellular degeneration takes place. This has been reported in Gossypium (Jensen & Fisher, 1969) and would appear to be a general feature of pollen tube growth, irrespective of its compatibility. Although results have yet to be analysed statistically, more transmitting tissue cells appear to degenerate in the vicinity of the growing pollen tubes than elsewhere in the tissue. The fate of the cytoplasm released into the intercellular space by the rupture of these protoplasts is not yet clear, but it may often be seen to invest the pollen tubes.

The metabolism of starch over the course of pollination

There is little doubt that a slight increase in stylar starch synthesis is stimulated by pollination itself. As with the previous cytoplasmic effects, it is not yet clear how the stimulus for this process is transmitted from the stigma. Other changes in the carbohydrate metabolism of the pistil also occur at this point, for Tupy (1961), working on Nicotiana, has reported an increase in glucose and fructose levels at the ovary 1 day after pollination, combined with a decline in the stigmatic sugars.

The pronounced drop in starch levels accompanying the growth of compatible pollen tubes is not unexpected. Starch is clearly an energy reserve, and if the tube is growing heterotrophically, depletion of such reserves should certainly occur. These observations are consistent with those of Linskens (1955) who reported free sugars to decrease to below half their pre-pollination levels following the passage of the pollen tubes, and of Roggen (1967) who demonstrated induction of enzymes for carbohydrate metabolism in the vicinity of the growing pollen tubes. The uses to which these extracellular carbohydrates may be put are many. O’Kelley (1955) describes their utilization in respiration, while Kessler, Feingold & Hassid (1960) have, in studies in vitro, described their employment in the synthesis of sucrose, callose and starch. In an investigation into the fate of the intracellular pectic substances, Kroh et al. (1970) and Kroh & Helsper (1974), reported their incorporation into the pollen tube wall.

The lack of starch mobilization following the passage of incompatible tubes is not so easily understood. This does not simply result from fewer tubes generally growing in such circumstances, for areas at the top of the transmitting tissue containing equivalent numbers of pollen tubes exhibit differences in starch metabolism dependent upon the compatibility of the cross. Again, biochemical investigations tend to support these observations; Linskens (1953, 1955) has reported a decrease in respiratory rate of pistils 12 h following self-pollination, and also differences between selfed and crossed styles in their endogenous levels of ‘glucan-hydrolases’ (Linskens et al. 1969). Taken in toto this evidence indicates that, in incompatible tubes, major pathways of carbohydrate metabolism are either blocked, or alternatively not activated.

The structural differences in the transmitting tissue following compatible or incompatible pollinations

Since the difference in pollen tube number normally encountered between compatible and incompatible crosses might cause effects that would be incorrectly interpreted, results are only discussed from the upper regions of the transmitting tissue where numbers of pollen tubes are equal irrespective of their compatibility and true stylar changes are thus most conspicuous.

The very different aspects displayed by transmitting tissue permeated by compatible tubes, and that containing incompatible tubes may almost be fully explained in terms of the utilization of reserves. The tissue with incompatible tubes looks very similar to that of pollinated flowers prior to the passage of the pollen tubes, while that in contact with compatible tubes is deficient in reserves in the form of starch and lipid, and contains far fewer microbodies.

Such a ‘degeneration’ of stylar tissue following the passage of compatible pollen tubes was reported by Crang (1966) to occur in Lychnis. In another species, Lilium, Crang (1969) found degeneration to occur in the parenchymatous cells investing the stylar canal cells, although these latter cells seemed not to be affected. While this work did not indicate clearly the cause of this degeneration, Crang (1969) proposed it to result from either enzymic secretion by the pollen tubes, or the stimulation by the pollen tube of ‘autolytic bodies’ in the stylar cells.

Although Rosen & Thomas (1970) confirmed that no ultrastructural changes overcame the stylar canal cells of Lilium on pollination, they reported an increase in secretory activity by these cells. Likewise, Yamada (1956) described the loss of cytoplasmic organization by the parenchymatous cells after pollination in Lilium, and pointed to the coincident arrival of a mucilagenous substance on the surface of the canal cells. The fact that starch and lipid were the first components of the cytoplasm to disappear from the cells of Lychnis and Lilium led Crang (1969) to propose that these reserves were utilized in the nutrition of the pollen tubes. In Gossypium, on the other hand, the position appears to differ, for Jensen & Fisher (1969) report the maintenance of stylar starch and lipid levels over the course of pollination. Results from the present investigation, however, concur well with those of Crang (1969), Yamada (1965), Rosen & Thomas (1970) and indicate that compatible pollen tubes stimulate the mobilization of stylar reserves, and subsequently utilize these products in the course of their metabolism.

Whilst this difference in metabolism of reserves between compatible and incompatible tubes is doubtless of significance, it is perhaps the events immediately prior to, and following pollination that are possibly most important to our understanding of the incompatibility system in Petunia. In an autoradiographic investigation, Labarca & Loewus (1973) reported that secretion by cells of the transmitting tissue into the intercellular space was independent of pollination, results supported by the electron-microscopic work of Sassen (1974) which indicated a release of intercellular substances throughout maturation of the style, reaching a maximum prior to anthesis. In a more recent investigation Cresti et al. (1976) have examined the formation of the intercellular matrix in Lycopersicum, and report that first secreted are pectic substances followed, after conspicuous activity of the endoplasmic reticulum and associated polyribosomes, by the formation of a small amount of protein.

While our results agree with this in part, there is little doubt that there are at least 3 phases of synthesis, one massive and prior to pollination, one stimulated by pollination and involving the ‘embayments’, and a final phase of secretion when mobilized stylar reserves are transferred to the pollen tube. The second phase occurs prior to the passage of the pollen tubes and is independent of compatibility, while the third occurs during pollen tube growth, and depends upon compatibility. These events may clearly be explained in terms of the model of van der Donk (1975) in which the S gene acts as a ‘master-gene’, switching on a battery of stylar genes that result in the support of pollen tube growth. In this connexion, it is noteworthy that little starch and lipid appears to be mobilized in transmitting tissue cells on self-pollination.

Alternatively, the differences observed between compatible and incompatible tubes may result from heterotrophic and autotrophic styles of growth. This concept is not new, for Rosen & Gawlick (1966) suggested, from work on Lilium, that changes they observed in incompatible tubes could be explained in terms of an inability to switch from an autotrophic to heterotrophic form of growth. Although this proposal has received much consideration in the literature (Kroes, 1973; Vasil, 1974), it has little support from the autoradiographic work of Kroh et al. (1970), where incompatible tubes were shown to take up more labelled precursor from the style than compatible tubes. This work, though far from conclusive, and the results of the study presented here would perhaps indicate that a metabolic deficiency lies within the incompatible tube itself.

Whatever model of pollen tube metabolism is finally adopted, the fact remains that incompatible pollen tubes do germinate, stimulate stylar metabolism and must utilize a portion, albeit very small, of the stylar reserves. At first sight this might appear to to be an inefficient use of stylar reserves, but under poor cross-pollination conditions such a mechanism enables self-pollination to ‘prime’ the style for the growth of the few cross-pollen tubes available.

The precise point of recognition presumably occurs when the pollen tube makes contact with S-specific polypeptides, or a larger molecular assembly containing them. The nature of these molecules and, indeed, their receptors in the pollen tube are not known. It is probably in species in which the contents of the stylar canal may be extracted without physiological damage to the surrounding cells, that the search for these recognition molecules may most profitably be carried out.

Thanks are due to the ARC and OECD for financial support, to the Royal Society for the provision of photomicrographic equipment, and to Ursula Potter for help with the illustrations.

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