AOBPreview originally published online on July 24, 2007
Annals of Botany 2007 100(3):459-470; doi:10.1093/aob/mcm137
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Seed Development in Ipomoea lacunosa (Convolvulaceae), with Particular Reference to Anatomy of the Water Gap
1 Department of Biology, University of Kentucky, Lexington, KY 40506, USA
2 Department of Horticulture
3 Department of Plant and Soil Sciences, University of Kentucky, Lexington, KY 40546, USA
* For correspondence. E-mail ccbask0{at}uky.edu
Received: 12 April 2007 Returned for revision: 2 May 2007 Accepted: 23 May 2007 Published electronically: 24 July 2007
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSLITERATURE CITED |
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Background and Aims: Disruption of one or both of the bulges (water gap) in the seed coat adjacent to the micropyle is responsible for breaking physical dormancy (PY) in seeds of Ipomoea lacunosa and other taxa of Convolvulaceae. Hitherto, neither ontogeny of these bulges nor onset of PY together with anatomical development and maturation drying of the seed had been studied in this family. The aims of this study were to monitor physiological and anatomical changes that occur during seed development in I. lacunosa, with particular reference to ontogeny of the water gap.
Methods: Developmental anatomy (ontogeny) of seed coat and dry mass, length, moisture content, germinability and onset of seed coat impermeability to water were monitored from pollination to seed maturity. Blocking/drying and dye-tracking experiments were done to identify site of moisture loss during the final stages of seed drying.
Key Results: Physiological maturity of seeds occurred 22 d after pollination (DAP), and 100 % of seeds germinated 24 DAP. Impermeability of the seed coat developed 27–30 DAP, when seed moisture content was 13 %. The hilar fissure was identified as the site of moisture loss during the final stages of seed drying. The entire seed coat developed from the two outermost layers of the integument. A transition zone, i.e. a weak margin where seed coat ruptures during dormancy break, formed between the bulge and hilar ring and seed coat away from the bulge. Sclereid cells in the transition zone were square, whereas they were elongated under the bulge.
Conclusions: Although the bulge and other areas of the seed coat have the same origin, these two cell layers underwent a different series of periclinal and anticlinal divisions during bulge development (beginning a few hours after pollination) than they did during development of the seed coat away from the bulge. Further, the boundary between the square sclereids in the transition zone and the elongated ones of the bulge delineate the edge of the water gap.
Key words: Convolvulaceae, hilar fissure and seed drying, Ipomoea, onset of seed coat impermeability, ontogeny of seed coat, physical dormancy, physiological maturity, water gap
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSLITERATURE CITED |
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Dormancy caused by a water-impermeable seed (or fruit) coat (i.e. physical dormancy, PY) is known to occur in 16 angiosperm families, but in no gymnosperms (Baskin et al., 2000, 2006). A palisade layer(s) in the seed (or fruit) coat is (are) responsible for this impermeability to water (Van Staden et al., 1989; Li et al., 1999a, b). Water enters the seed when the ‘water gap’, a special morpho-anatomical area in the seed coat, opens. Ecologically, the water gap serves as an environmental signal detector for dormancy break to occur when conditions are suitable for seed germination and plant survival (Baskin and Baskin, 2000).
The onset of water impermeability during development of seeds with PY is well documented in Fabaceae [Trifolium pratense, T. repens, Lupinus arboreus (Hyde, 1954), Vicia villosa (Jones, 1928) and Cassia acutifolia (Bhatia et al., 1977)], Anacardiaceae [Rhus aromatica and Rhus glabra (Li et al., 1999c)] and Malvaceae [Sida spinosa (Egley, 1976) and Gossypium hirsutum (Patil and Andrews, 1985)]. However, development of physical dormancy in Convolvulaceae has not been described accurately.
Development of the seed coat in seeds with PY has attracted the attention of scientists for many years, but most studies have not followed a timeline of development, such as days after pollination (DAP). Instead, ovules and various stages of seed development have been studied. Development of the water gap in seeds of Fabaceae, i.e. lens (sometimes misleadingly called strophiole) was documented for Indigofera parviflora by Manning and Van Staden (1987) and for Trifolium repens by Martens et al. (1995). Both of these studies followed development of the lens and of the seed coat away from the lens from ovule to mature seed without any timeline. Development of the water gap in Cannaceae (Canna tuerckeimii) was well documented on a timeline from floral initiation to mature seed by Grootjen and Bouman (1988). In Malvaceae, water-gap development was studied in Sida spinosa by Egley and Paul (1982) and in Abutilon theophrasti by Winter (1960) from ovule to mature seed following a timeline. Chopra and Kaur (1965) studied seed development in Bixa orellana (Bixaceae); however, they described the anatomy of the water gap, i.e. chalazal cap, only in the mature seed. Li et al. (1999a, b) described ontogeny of the water impermeable endocarp and water gap, i.e. carpellary micropyle, in Rhus aromatica and R. glabra (Anacardiaceae) using seeds (endocarp + true seed) randomly collected in various stages of development.
Water-gap development in the other ten families in which PY is known to occur, including Convolvulaceae, has not been documented. However, seed-coat and hilar-pad development have been described for several species in this family: Ipomoea rubra-caerulea (= I. violacea) (Woodcock, 1942), Convolvulvus arvensis (Sripleng and Smith, 1960), I. sinuata (= Merremia dissecta), I. purpurea and I. carnea (Kuar and Singh, 1970), I. purpurea and Rivea hypocrateriformis (Govil, 1970) and 33 other species (Kuar and Singh, 1987). However, none of them described development of the water gap. Chandler et al. (1977) monitored changes in dry and wet seed mass and of germinability of I. turbinata from ovule to mature seed, but they did not describe seed coat anatomy.
The objectives of the present study were 3-fold: (1) to monitor physiological changes that occur during seed development in Ipomoea lacunosa; (2) to describe maturation drying of seeds of this species; and (3) to describe the developmental anatomy of the water gap. Ipomoea lacunosa belongs to Ipomoeeae, the most advanced tribe in Convolvulaceae (Stefanovic et al., 2003), and it is native to the eastern USA (Abel and Austin, 1981; Elmore et al., 1990; Gleason and Cronquist, 1991). This species is a summer annual vine (Gleason and Cronquist, 1991) that is a bad weed in corn, cotton (Anon., 1995), soybeans (Anon., 2000) and rice (Anon., 2001). Seeds of I. lacunosa are physically dormant at maturity, and dormancy can be broken by mechanical scarification (Egley and Chandler, 1978), via opening of the water gap, by dipping seeds in boiling water for 4 s, or by incubating them at 35 °C for 
3 h at high relative humidity (RH) (Jayasuriya et al., 2007). When seeds become non-dormant, a slit is formed around one or both of the bulges of the seed coat located adjacent to the hilum and micropyle. Each bulge is a specialized anatomical structure with a transition area between it and the seed coat away from the bulge (Jayasuriya et al., 2007).
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSLITERATURE CITED |
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Growing plants
Plants of I. lacunosa (Gunn, 1969; Jones, 2005; D. F. Austin, Florida Atlantic University, pers. comm.) were grown from seeds collected in a corn field at Spindletop Farm, University of Kentucky, Lexington, KY, USA, in October 2005. The seeds were manually scarified and planted in pots containing greenhouse potting soil in a non-heated greenhouse on the University of Kentucky campus in Lexington in May 2006. Plants were watered as required, and insect pests were controlled using insecticides.
Pollination
Flowers were pollinated between 0530 h and 0630 h, before they opened. Pollination was done by touching mature stigmas with anthers releasing mature pollen. After pollination, flowers were covered with vanilla paper bags. Flowers were tagged to keep track of their development. Bags were removed from the flowers 5 d after pollination to allow fruits to grow naturally.
Seed development and moisture content
Developing fruits and seeds were collected on various days after pollination (DAP; see Fig. 1). Lengths of 30 seeds and of 30 fruits were measured using a calibrated dissecting microscope. Six replicates of 15 seeds were weighed to the nearest 0·0001 g, and seeds were oven-dried to constant mass at 105 °C. Seed moisture content was determined on a fresh mass basis.
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The mass of 15 seeds each was determined at time 0, and seeds were placed on moist filter paper in Petri dishes. At 1-h intervals, until all seeds had fully imbibed, they were removed from the Petri dish, blotted dry with filter paper, weighed to the nearest 0·0001 g and returned to the wet filter paper in the dish. Moisture content was determined based on initial fresh mass basis.
Germinability of developing seeds
Three replicates of 15 seeds for each collection date were placed on moist sand in Petri dishes and incubated at 25/15 °C (12/12 h) in light/dark (14/10 h; approx. 40 µmol m–2 s–1, 400–700 nm, cool white fluorescent light). When seeds became impermeable (at 30 DAP), both manually scarified and non-scarified seeds were tested for germination.
Site of water loss during maturation drying
Blocking experiments
Seeds were collected 25 DAP and grouped into four colour categories – white, brown, dark-brown and black – which represented increased levels of maturation. The hilum, bulge and bulge plus hilum of 15 seeds in each category were painted with Thompson's water seal (Thompson and Formby, Inc., Memphis, TN, USA). No paint was applied to 15 control seeds in each of the four maturation stages. Seeds in each category were weighed separately and allowed to air-dry, in separate Petri dishes with the tops removed, at ambient laboratory conditions (22–24 °C, 55–60 % RH). Seeds were weighed at time 0 and at 1-h intervals for 12 h and then daily until there was no further decrease in mass. Then, they were oven-dried to constant mass at 105 ºC.
Dye-tracking experiments
Seeds were collected 25 DAP and grouped into the four colour categories described above. Twenty seeds from each category were placed separately in aniline-blue solution. Two seeds from each category were removed at 15-min intervals for 150 min. Longitudinal and transverse cuts were made of these seeds and the staining pattern observed and recorded.
Development of seed coat and of water gap
Developing seeds were collected at 0, 1, 2, 3, 4, 6, 8, 10, 12, 15, 20, 22, 25, 28 and 30 DAP. Seeds were fixed in FAA (formaldehyde : glacial acetic acid : ethyl alcohol : water) solution (2 : 1 : 10 : 7, v/v/v/v), dehydrated using the ethanol–TBA series and embedded in paraffin (paraplast plus). Sections (16 µm thick) were made with a hand rotary microtome (Leica RM 2135). Paraffin was removed using the Hemo-De ethanol gradient, and tissue was stained with 3 % safranin in water and then with 1 % fast green in 95 % ethanol. Sections were observed with an Olympus light microscope, and photographs were taken of the bulge and hilum area with a Canon EOS 30 D camera. Photographs were used to trace development of the bulge.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSLITERATURE CITED |
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Seed development and moisture content
Seeds attained maximum dry mass (i.e. physiological maturity) about 22 DAP (Fig. 1). Moisture content of developing seeds increased from about 70 % at day 0 to about 85 % at 10 DAP, after which it decreased gradually to about 65 % at 26 DAP. Between 26 and 32 DAP, seeds dried rapidly to a moisture content of 13 %. Fruit length and seed length increased rapidly for 15 d and did not change thereafter. Reduction of seed length followed water loss (Fig. 2).
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Seed moisture content was correlated with the seed coat impermeability. When seed moisture content had decreased to
13 %, seed coats of all seeds had become impermeable to water (Fig. 3).
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Germinability of developing seeds
Increase in dry mass was sigmoidal, reaching an asymptote at 22 DAP (Fig. 4). Germinability of a portion of the seeds was attained by 20 DAP, and by 24 DAP 100 % of the seeds could germinate. Seeds had become impermeable by 30 DAP and did not germinate unless they were manually scarified.
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Site of water loss during maturation drying
Blocking experiments
Painted and non-painted seeds at the white stage of development lost the same amount of water (Fig. 5A). However, painting the hilum of seeds at the brown, dark brown and black stages significantly reduced water vapour loss (Fig. 5B–D). Water loss in seeds having the bulge painted did not differ from those in which the bulge was not painted (control).
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Dye-tracking experiments
After seeds were soaked in aniline-blue dye for 15 min, the entire seed coat of white seeds was stained, whereas only the cells under the hilum were stained in brown, dark brown and black seeds (data not shown).
Development of seed coat and of water gap
Ipomoea lacunosa has a unitegmic, anatropous ovule (Corner, 1976). The integument cannot be separated clearly from the nucellus; however, there are two distinct uniseriate layers in the ovule that give rise to the seed coat in the mature seed (Fig. 6.). These two layers are the outermost ones in the ovule (Figs 7A, B, 8A and 9A). They consist of square-shaped cells with large nuclei. Staining of these nuclei with safranin confirmed that they are capable of meristematic activity.
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Development of the seed coat
The ontogenetic sequence of development of the seed coat away from the bulge (hereafter seed coat) is shown in Fig. 6. Development of cell layers of the seed coat were completed within 12 DAP (Figs 6 and 8D). However, cell wall thickening and chemical changes in cell walls occurred until 30 DAP. Periclinal division of the outermost layer of the integument gave rise to two layers by 1 DAP. The outer layer of these two new layers consisted of large bulging epidermal cells (‘bulging cell layer’ sensu Kuar and Singh, 1987), most of which fell off the seed when it became mature. The second layer, which consisted of flattened rectangular-shaped cells, gave rise to the epidermis. Periclinal division of the second layer of the integument occurred within 5–6 DAP (Fig. 8B). Cells of the two layers produced by these periclinal divisions elongated and formed two elongated hypodermal cell layers by 8 DAP (Fig. 8C). These cell layers contained large nuclei and seemed to be meristematic at 8 DAP. Anticlinal divisions of the outer elongated hypodermal layer resulted in the formation of thin palisade-like cells, while periclinal divisions of the innermost layer produced 4–5 square- to polygonal-shaped cell layers by 10 DAP (Fig 6). By 12 DAP (Fig. 8D), the walls of palisade layer cells had become thicker and the cells were dead each one containing a clear small lumen. By 15 DAP, four cell layers below the palisade layer had given rise to sclereids (Fig. 8E). The shape of these cells in the seed coat and in the bulge–seed coat transition area is square. However, their shape gradually changes from square to elongate in areas away from the hilum and bulge (Fig. 8F).
Development of the bulge
The ontogenetic sequence of bulge formation is shown in Fig. 6. The bulge is morphologically and anatomically distinct from the rest of the seed coat, even in ovules 2–3 h after pollination. In the bulge area, there are three distinct uniseriate layers compared with two in the seed coat (Fig. 9A). However, the one outermost layer and the two innermost layers seem to originate from two separate layers. Similar to what happened in the development of the seed coat, the outermost layer gave rise to a large bulged-epidermal cell layer that fell off with later development of the seed coat. The adjacent layer gave rise to the epidermis, which consisted of flattened rectangular-shaped cells. By 1 DAP, three hypodermal layers had been produced by periclinal divisions of the two innermost layers of the integument.
By 2 DAP, the outermost hypodermal layer had undergone further periclinal divisions to produce one additional layer, whereas the rest of the hypodermal layers had undergone anticlinal divisions only (Fig. 9B). These four hypodermal layers consisted of square-shaped cells with large nuclei and were meristematic. The newly formed outer hypodermal layers produced four new cell layers by further periclinal divisions. Of these four cell layers, the outermost ones underwent more anticlinal divisions and elongation and formed two elongated layers. Therefore, at 4 DAP the hypodermal layer consisted of two elongated and four square-shaped cell layers (Fig. 9C). The outer square-shaped cell layers produced two more cell layers, and these four layers had elongated by 5 DAP.
At 5 DAP, the endodermal layer consists of six elongated cell layers. Between 5 and 10 DAP (Fig. 9D), the outermost elongated layer underwent anticlinal divisions and gave rise to a palisade layer. Cell walls of the palisade layer under the bulge became thickened, and cells died before they did so in the seed coat away from the hilum and bulge. At 10 DAP, all of the cell layers were present in the bulge. Thus, the bulge consisted of remnants of the bulging epidermal cells, a small flattened dead epidermis, a palisade layer, six elongated cell layers and two to four square-shaped cell layers (Fig. 10). Transition of elongated cells to square cells between the bulge and seed coat away from the hilum and bulge, and of elongated cells to square and polygonal cells between the bulge and hilar area, was evident by 10 DAP (Fig. 9E). After 10 DAP, the shape of the cells in the bulge did not change; however, cell walls of these cells became thicker and the size of the cell lumen decreased gradually (Fig. 9F–H).
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Development of the hilar pad
The ontogenetic sequence of hilar pad development is shown in Fig. 6. The hilar region is a distinct anatomical area in the mature seed coat. Even in the ovule it is distinct from the other two areas, i.e. bulge and seed coat away from the bulge. The funiculus is attached to the hilar fissure that surrounds the hilar pad, and there is no physical connection between funiculus and hilar pad (Fig. 7A–C). Therefore, the funiculus forms a cup-like structure around the hilar pad. The hilar pad consists of a distinct uniseriate layer that develops into the seed coat in the hilar pad (Fig. 10A). The vascular system enters the seed through the funiculus/hilar fissure connection. It entered the seed all around the hilar pad, connected below the hilum and then divided into two branches. One branch extended to near the embryo sac, whereas the other one extended downward and curved at the base of the seed. This branch extends all the way to the micropyle (Fig. 7B, C).
The cell layer in the hilar pad divided periclinally and had formed four cell layers 1 DAP. The outermost of these four layers divided periclinally to form one other cell layer between 3 DAP and 4 DAP (Fig. 10C). Thus, at 4 DAP the hilar pad consisted of five cell layers. All five layers have square-shaped cells with large nuclei. The innermost layer had elongated by 5 DAP (Fig. 10D). By 8 DAP, this elongated layer had divided periclinally and formed another elongated layer. The two innermost layers of square-shaped cells also had elongated at this time. Therefore, at 8 DAP the hilar pad consisted of two layers of square-shaped cells and four layers of elongated cells. The innermost elongated cell layer divided anticlinally and formed narrow elongated cells, which later developed into the counter palisade. Cells in this layer contained globular nuclei at their centre. The second layer of elongated cells divided periclinally again and formed another layer of elongated cells. The third layer of elongated cells divided anticlinally to form the palisade layer. At 10 DAP, cells of the palisade layer were thickened and dead. The fourth layer of elongated cells underwent further periclinal divisions and formed two to four square-shaped cells. At 12 DAP, the hilar pad consisted of two outermost layers of square cells, a layer of elongated cells that give rise to the counter palisade, two layers of elongated cells, a palisade layer and two to four layers of square-shaped cells (Figs 6 and 10E). These are the cell layers in the mature seed coat. After 12 DAP, the shape of cells in the hilar pad did not change; however, cell walls of these cells became thicker, and the size of the cell lumen decreased gradually (Fig. 10F–H).
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DISCUSSION |
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As expected, the growth curve for I. lacunosa seeds was sigmoidal. Seeds were physiologically mature at 22 DAP, and thus they did not accumulate additional dry matter after this time. At the beginning of seed development, the water content of I. lacunosa seeds increased, as reported for developing seeds of other species (Chandler et al., 1977; Welbaum and Bradford, 1988; Hong and Ellis, 1990; Li et al., 1999c). Maturation drying began after seeds attained physiological maturity. During maturation drying, I. lacunosa seeds lost water vapour rapidly. Within 5 d, moisture content dropped from 65 % to 13 %. This rapid drying has also been observed in the development of physically dormant seeds in species of Fabaceae (Hyde, 1954), Malvaceae (Patil and Andrews, 1985) and Anacardiaceae (Li et al., 1999c). During maturation drying, seed colour changed from white to black, with the change in colour beginning in the hilar–micropylar area. During this time, cell walls of cells in the palisade layer thickened, and cell wall thickening began in cells in the micropylar area. Palisade cells of the seed coat became thick 2–3 d later than those in the micropylar area did. The seed coat gradually and sequentially became impermeable during maturation drying, as seed colour changed from white to brown to dark brown and finally to black.
Dye-tracking experiments showed that the seed coat is completely impermeable in brown seeds, in which the palisade cells are thickened and dead throughout the seed coat. However, the hilar fissure is open in brown seeds, and this is the place through which final drying takes place (Fig. 5). Dye-tracking experiments (data not shown) provide further support for this conclusion. Observations on maturation drying in seeds of Fabaceae species with physical dormancy (Hyde, 1954; Quinlivan, 1971) are in agreement with those on I. lacunosa. For Cercis canadensis (Fabaceae), Jones and Geneve (1995) advanced the idea that the hilar slit might be both the last place of water loss during seed drying and the initial place of water entry into seeds during imbibition; however, this is not the case for Convolvulaceae. The present experiments clearly show that the slits around the bulges are the initial routes of water entry into seeds of I. lacunosa (Jayasuriya et al., 2007), while the hilar fissure is the last place to become impermeable to loss of water vapour from the seed during maturation drying (Fig. 8).
When seeds dried to 13 % moisture content, the hilum fissure closed tightly, and thus the entire seed coat became impermeable. Therefore, seeds placed on wet filter paper 30 DAP did not imbibe water (Fig. 2), and thus they did not germinate (data not shown). This has been observed in physically dormant seeds of Fabaceae (Hyde, 1954); however, the moisture content at which seeds become impermeable differs among species. Ipomoea turbinata seeds became impermeable at 8·5 % (Chandler et al., 1977), and those of I. pes-tigridis dried at 60 °C became impermeable at 2·5–10 % (Bhati and Sen, 1978). The moisture content at which seeds in other families attain impermeability also varies, e.g. in Fabaceae: Crotalaria spectabilis, 11 % (Egley, 1979); Gymnocladus dioica, 5–8 % (Raleigh, 1930); Lupinus arboreus, 11 % (Hyde, 1954); Sesbania bispinosa, 9·6 % (Graff and Van Staden, 1987); Trifolium pratense and T. repens, 14 % (Hyde, 1954); and Vicia villosa, 14 % (Jones, 1928); in Malvaceae: Gossypium hirsutum, 12 % (Patil and Andrews, 1985) and Sida spinosa (Egley, 1976); and in Anacardiaceae: Rhus aromatica, 6·1 % and Rhus glabra, 9·3 % (Li et al., 1999a).
Germinability of I. lacunosa seeds was 100 % at 24 DAP, which was 2 d after physiological maturity. Seeds of I. turbinata (Chandler et al., 1977) germinated to 100 % at physiological maturity. However, most crop seeds attain the ability to germinate to 100 % well before physiological maturity (TeKrony and Egli, 1995).
Like other species of Convolvulaceae, I. lacunosa has an anatropous ovule (Corner, 1976). In I. lacunosa, the integument and nucellus of the ovule cannot be distinguished from each other, in agreement with observations of Woodcock (1942) for I. rubro-caerulea (= I. violacea). However, the integument of I. sinuata (= Merremia dissecta), I. purpurea, I. carnea (Kaur and Singh, 1970), Convolvulus arvensis (Sripleng and Smith, 1960) and several other species (Kaur and Singh, 1987) is 15–25 cells thick. The integument of I. lacunosa is 15–20 cells thick according to our estimation based on the position of the vascular bundle. However, in several species of Ipomoea (Woodcock, 1942; Kaur and Singh, 1970, 1987) and in Convolvulvus arvensis (Sripleng and Smith, 1960) only the two outermost layers of the integument participate in seed-coat development. According to Kaur and Singh (1970, 1987) and Sripleng and Smith (1960), the second outermost layer in the mature seed coat develops from the hypodermal layer in several Convolvulaceae species.
Thus, it is clear in I. lacunosa that the second outermost layer, which is called the epidermis in this paper, develops by periclinal division of the dermatogen, which is the outermost layer in the ovule. The outermost layer of the integument in I. lacunosa and in other Convolvulaceae species (Woodcock, 1942; Sripleng and Smith, 1960; Kaur and Singh, 1970, 1987), which develops from the dermatogen, forms a layer of bulging epidermal cells that may develop into hairs in some Convolvulaceae species, e.g. I. batatas, I. crassicaulis (= I. carnea), I. maxima, I. obscura, I. pes-caprae, Merremia hederacea (Sampathkumar and Ayyangar, 1982), I. purpurea and I. carnea (Kaur and Singh, 1970). As in Convolvulus arvensis (Sripleng and Smith, 1960), I. rubro-caerulea (= I. violacea) (Woodcock, 1942) and several other Convolvulaceae species (Kaur and Singh, 1970, 1987), the palisade layer of the seed coat in I. lacunosa develops from the hypodermal layer. Compared with the single palisade layer in the mature seed coat of I. lacunosa, there may be two palisade cell layers in the mature seed coat of Calystegia sepium, Cuscuta chinensis, I. sepium (= Calystegia sepium), I. hederacea, I. leari, I. purpurea, I. fistulosa, Quamoclit coccinea (probably I. hederifolia sensu D. Austin, Florida Atlantic University, pers. comm.) (Hamed and Mourad, 1994), I. angulata (= I. hederifolia), I. coccinea, I. hispida (Kaur and Singh, 1987), I. carnea, I. sinuata (= Merremia dissecta) and I. purpurea (Kaur and Singh, 1970), two to three in the mature seed coat of I. ramoni (= I. triloba) (Kaur and Singh, 1987) and three in the mature seed coat of I. angustifolia (= Xenostegia tridentata), I. tenuipes and Mina lobata (= I. lobata) (Kaur and Singh, 1987).
As in C. arvensis (Sripleng and Smith, 1960), the hilar pad of I. lacunosa develops from a single cell layer. In contrast, the seed coat in this species develops from two cell layers. Instead of giving rise to an epidermis and to a layer of bulging epidermal cells, the innermost layer of the hilar pad underwent a series of periclinal divisions and gave rise to a thick pad. However, the palisade layer of the hilar pad area developed from the first outermost layer below the dermatogen. This development of the palisade layer is similar to that of the seed coat away from the bulge. Although many people have studied the ontogeny of Convolvulaceae seeds, including that of those of several Ipomoea species, they have not described the differences in the way the funiculus is attached to the seed. In most seeds, the funiculus is attached directly to the hilar pad. In I. lacunosa, on the other hand, the funiculus is attached to the hilar pad only through the hilar fissure, which forms a ring around the hilar pad. Cells in the hilar pad have no physical contact with the funiculus.
Although the bulge is another distinct anatomical area of the seed coat, its ontogeny hitherto has not been studied. However, the development of the water gap has been studied in several other families known to have PY, including the chalazal plug in several families of Malvales [Malvaceae (Winter, 1960; Egley et al., 1986), Cistaceae (Thanos and Georghiou, 1988), Cochlospermaceae, Dipterocarpaceae and Sarcolaenaceae (Nandi, 1998)], the protuberance in Nelumbonaceae (Ohga, 1926), the so-called carpellary micropyle in Anacardiaceae (Li et al., 1999b), the operculum in Cannaceae (Grootjen and Bouman, 1988) and the lens in Fabaceae (Manning and Van Staden, 1987; Martens et al., 1995). The development of the bixoid chalazal plug in Malvales (Chopra and Kaur, 1965; Nandi, 1998) is considerably more complex than that of the water gap in Convolvulaceae.
Development of the lens in Fabaceae has some similarities to that of the bulge in Convolvulaceae. The bulge in Convolvulaceae and the lens in Fabaceae develop from the same ontogenetic layers that give rise to the seed coat away from the hilar region. However, change in the developmental series produces the anatomical and morphological uniqueness of the structure of the water gap in these two families. These structures in both families are swellings in the regular seed coat. In some legumes, palisade cells in the lens are taller (Manning and Van Staden, 1987) or shorter (Serrato-Valenti et al., 1995) than those in the seed coat away from the lens, whereas the palisade layer of the bulge in I. lacunosa is the same length as that in the seed coat.
The bulge of I. lacunosa develops completely from the two outermost cell layers of the integument, which are actively involved in development of the entire seed coat. Although the bulge and other areas of the seed coat have the same origin, these two cell layers underwent a different series of periclinal and anticlinal divisions in bulge development (beginning a few hours after pollination) than they did in the seed coat away from the bulge. Further, cell wall thickening in the bulge was faster than it was in the seed coat, which led to a swelling in that area of the seed coat, i.e. bulge formation.
As a result of this differential ontogenetical series, a transition zone formed between bulge and hilar ring and between bulge and seed coat away from the bulge. Especially in the bulge–hilar ring margin, sclereid cells change in shape from square to elongated (Fig. 9G, H). Further, a clear change in orientation of the palisade layer was seen between the bulge and the hilar ring. Moreover, sclereid cells change from elongated to square at the transition between bulge and seed coat adjacent to the bulge. These transition zones are weak margins where the seed coat ruptures, between bulge and hilum and between bulge and regular seed coat. These weak margins seemed to rupture due to a pressure developed in the hypodermal cells below the bulge. Water vapour is important in the formation of these slits when seeds are not responding to dry heat (Jayasuriya et al., unpubl. res.). We speculate that hydration of hypodermal cells due to water vapour that enters into seed through micro-openings in the hilum, coupled with high temperature, causes swelling of cells in the hypodermal layer. This may be the source of pressure that causes formation of the slit around the bulge. Therefore, this special anatomical transition appears to be important in the response of seeds to the environmental cue that causes the slit to be formed, i.e. the dormancy break, after which the seed imbibes water and germinates.
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We thank Dr Sharyn E. Perry, Department of Plant and Soil Sciences, University Kentucky, for use of microtome and tissue-sectioning equipment; Ms Sharon T. Kester, Department of Horticulture, University of Kentucky, for help with sectioning and staining of tissues; and Dr Daniel F. Austin, Florida Atlantic University, for help with the taxonomy and nomenclature of Ipomoea.
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