Annals of Botany 92: 571-580, 2003
© 2003 Annals of Botany Company
Systematic Significance of Cell Inclusions in Haemodoraceae and Allied Families: Silica Bodies and Tapetal Raphides
1 Royal Botanic Gardens, Kew, Richmond, Surrey TW9 3DS, UK
* For correspondence. E-mail c.prychid@kew.org
Received: 10 January 2003;; Returned for revision 15 April 2003. Accepted: 13 June 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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This paper presents the first record of silica deposits in tissues of Haemodoraceae and adds new records of tapetal raphides in this family. Within the order Commelinales, silica is present in leaves of three families (Hanguanacaeae, Haemodoraceae and Commelinaceae), but entirely absent from the other two (Pontederiaceae and Philydraceae). Presence or absence of characteristic cell inclusions may have systematic potential in commelinid monocotyledons, although the existing topology indicates de novo gains and losses in individual families. Silica sand was observed in leaves of five out of nine genera examined of Haemodoraceae, predominantly in vascular bundle sheath cells and epidermal cells. Within Haemodoraceae, silica is limited to subfamily Conostylidoideae. The occurrence of silica in Phlebocarya supports an earlier transfer of this genus from Haemodoroideae to Conostylidoideae. The presence of raphides (calcium oxalate crystals) in the anther tapetum represents a rare character, only reported in a few monocot families of the order Commelinales, and possibly representing a mechanism for regulation of cytoplasmic free calcium levels. Tapetal raphides were observed here in Anigozanthus and Conostylis (both Haemodoraceae), and Tradescantia (Commelinaceae), thus supplementing two earlier records in Haemodoraceae, Philydraceae and Commelinaceae.
Key words: Silica sand, tapetal raphides, monocotyledons, Haemodoraceae, Commelinales.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Characteristic cell inclusions, especially different types of calcium oxalate crystals and silica bodies, often represent significant taxonomic characters at various taxonomic levels in flowering plants. For example, in monocotyledons, Prychid and Rudall (1999) demonstrated that the presence of styloids (solitary prismatic crystals) is potentially a highly consistent synapomorphy for some families (e.g. Iridaceae) or groups of families, and that druses (cluster crystals) are almost entirely restricted to basal monocots (Acorus, some Araceae and Tofieldia). Raphides (bundles of needle-like crystals) are widespread, but in monocots their absence is synapomorphic for some groups; e.g. raphides have apparently been ‘lost’ twice within the order Liliales (Rudall et al., 2000).
This paper reviews the occurrence and distribution of cell inclusions in the monocot family Haemodoraceae and other members of the order Commelinales in a systematic context. Haemodoraceae are a small family centred in south-western Australia (Macfarlane et al., 1987; Simpson, 1990, 1998) with two distinct subfamilies: Conostylidoideae (six genera) and Haemodoroideae (eight genera). There are four families of Commelinales sensu Angiosperm Phylogeny Group (1998): Haemodoraceae (14 genera), Commelinaceae (41 genera), Philydraceae (four genera) and Pontederiaceae (nine genera) (Kubitzki, 1998). Recent molecular evidence (e.g. Chase et al., 2000; Angiosperm Phylogeny Group II, 2003; Graham et al. 2003) has indicated that Hanguanaceae (which includes a single genus, Hanguana) may also belong in this group, probably as sister to Commelinaceae, although this placement is partially contradicted by morphological data, which indicate a closer relationship with Zingiberales (Rudall et al., 1999).
Commelinales are particularly variable for all types of cell inclusions and, therefore, merit further investigation for these characters, which may help to elucidate inter familial relationships. Hitherto, silica bodies have been recorded in only Commelinaceae and Hanguanaceae (Tomlinson, 1969), and not in the other three families of Commelinales. However, during the course of a current review of silica bodies in monocots (Prychid et al., 2003), silica was observed in several genera of Haemodoraceae, indicating that further exploration of this character within Commelinales may be worthwhile, especially in Haemodoraceae. A recent observation by Hardy and Stevenson (2000) of tapetal raphides in Commelinaceae also prompted further investigation of this relatively infrequent character, which has also been reported in the related family Philydraceae (Hamann, 1966).
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MATERIALS AND METHODS |
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Material examined
Leaves of the following species were examined for the presence of cell inclusions (HK indicates material cultivated at Kew, followed by Kew accession number; s.n. indicates data not available).
Silica.
Observations were made on leaves of 36 species of Haemodoraceae, representing nine out of 14 genera. In addition, microscope slides (in the collection at RBG Kew) of leaves of several species of Pontederiaceae, Philydraceae and Commelinaceae were examined for comparison (not listed here). Observations on silica bodies in other monocots are summarized by Prychid et al. (2003).
Conostylidoideae: Anigozanthos flavida DC. (HK 1952; HK 1983-4125), A. humilis Lindl. (s.n., Australia), A. manglesii D.Don. (HK 1952; s.n., Australia), A. pulcherrima Hook. (Morrison 15138, Australia), A. rufa Labill. (Morrison s.n., Australia), A. rufus (HK 1984-4484), A. viridis Endl. (s.n., Australia), Blancoa canescens Lindl. (Morrison 16156, Australia), Conostylis aculeata R.Br. (Green 1732, Australia), C. androstemma F.Muell. (Green E196, Australia), C. bealiana F.Muell. (Churchill and Green 1956, Australia), C. bracteata Endl. (Morrison 10197, Australia), C. breviscapa R.Br. (Churchill and Green 1956, Australia), C. bromelioides Endl. (s.n., Australia), C. candicans (Green 497.8/56, Australia), C. filifolia F.Muell. (Morrison 20086, Australia; Green E168, Australia), C. juncea Endl. (Helms 1899, Australia; Williams 9/68, Australia; Green E129, Australia), C. serrulata R.Br. (Green 1739, Australia), C. setigera R.Br. (s.n., Australia; Morrison 90, Australia; Green E143, Australia), C. setosa Lindl. (s.n., Australia), C. stylidoides F.Muell. (Green E149, Australia), C. vaginata Endl. (Grewar 1968, Australia), Macropidia fumosa J.Drumm. ex Harv. (Morrison 16161 and 13283, Australia; Williams 1968, Australia), Phlebocarya ciliata R.Br. (Morrison 118 1901, Australia; Williams, HK 1968), P. filifolia F.Muell. (Drummond 368 1844, Australia; Williams, HK 1968), P. pilosissima F.Muell. (Mueller 1873, Australia n.s.; Williams 1968, HK), Tribonanthes brachypetala Lindl. (Green E173, Australia; Williams 1968, Australia; Morrison 1899 No. 9266, Australia).
Haemodoroideae: Dilatris corymbosa Berg. (Parker s.n., South Africa; Williams s.n., 1968, HK), D. ixioides Lam. (Schlieben & van Breda, South Africa; Williams 9/’68, South Africa; Acocks 19837 x158, South Africa), D. pillansii W.F. Barker (Lenyns 1958, South Africa), Haemodorum coccineum R.Br. (Veitch 1866, Australia; Williams 1968, Australia), H. distichophyllum Hook. (Williams s.n., HK 1968; Long, Australia 1929), H. laxum R.Br. (Morrison 1897 No. 7238, Australia), H. simulans F.Muell. (Drummond 1839, Australia; Williams 1968, HK), H. spicatum R.Br. (Morrison 1904 No. 548, Australia; Williams 1968, Australia), H. teretifolium R.Br. (Bot. Gdns. Sydney Australia, 1952; Rodway 1936, Australia) dLachnanthes tinctoria Ell. (Biltmore Herb. No. 27288, Fayetteville, N. Carolina, USA), Wachendorfia brachyandra (W.F. Barker, South Africa 1957), W. thyrsiflora Linn. (Williams 1925–6, South Africa; Williams, HK 1968).
Tapetal raphides.
Flowers or anthers of the following species were examined for the presence of tapetal raphides.
Commelinaceae: Tradescantia ohiensis Rafin. (HK 1983-426), T. spathacea Sw. (HK 1992-2952).
Haemodoraceae: Anigozanthos (A. bicolor x A. humilis) x A. flavidus (HK 1991-891), A. flavidus DC. (HK 1983-4125), Conostylis candicans x aculeata (HK 1997-6468), Tribonanthes longipetala Lindl. (P.J. Rudall 20, Australia), Wachendorfia paniculata Linn. (HK 1988-3965).
Philydraceae: Helmholtzia glaberrima Caruel. (HK 1992-1642).
Pontederiaceae: Eichhornia azurea Kunth (HK 1975-1464), E. paniculata Solms (HK 1963-1714), Heteranthera callaefolia Reichb. ex Kunth (Nigeria, R.D. Meikle, No. 863), Monochoria africana N.E.Br. (Kenya, P.J. Greenway, No. 9483), Pontederia cordata var. cordata (HK 1969-50051), P. lanceolata Nutt. (HK 1969-50052).
Methods
Leaves were sectioned on a sliding microtome and stained with safranin and Alcian Blue before being dehydrated through an alcohol series, taken through Histoclear and mounted onto microscope slides in Euparal.
Flowers and leaves were fixed in formalin–acetic–alcohol (FAA). Material was prepared for scanning electron microscopy (SEM) by dehydration through an alcohol series, critical point drying using a Bal-Tec 030 critical point dryer, dissection onto pin stubs, sputter coating with platinum and examination using a Hitachi S4700 FE-SEM. Material was prepared for light microscopy (LM) using standard methods of paraffin embedding (Johansen, 1940) and sectioned using a rotary microtome. Serial sections, 14 µm thick, were stained with safranin and Alcian Blue, dehydrated through an ethanol series to 100 % ethanol, and transferred to Histoclear before being mounted in DPX.
For examination of Tradescantia ohiensis and Anigo 0zanthos flavidus with both LM and transmission electron microscopy (TEM), fresh buds in a range of sizes were selected for each species. Anthers were dissected out and placed in 2·5 % glutaraldehyde in 0·1 M cacodylate buffer (pH 7·2), de-aerated under vacuum for 1 h and fixed for 16–20 h at 4 °C. They were washed in cacodylate buffer, post-fixed in 1 % buffered osmium tetroxide for 3 h at room temperature, washed again, and dehydrated through a graded ethanol series. Anthers were then dehydrated in three changes of 100 % acetone and embedded in Spurr resin (Spurr, 1969) in flat moulds. Semi-thin (approx. 1 µm) sections were cut using a Reichert Ultracut and a dry glass knife, stained with toluidine blue and mounted in DPX (Aldrich). They were examined using a Nikon Labophot light microscope with camera attachment, using normal bright-field optics. Ultra-thin (gold) sections were then cut using a diamond knife, stained with uranyl acetate and lead citrate in an LKB Ultrostainer and examined using a JEOL JEM-1210 transmission electron microscope.
Identification of cell inclusions was based on shape and refraction characteristics. For instance, silica identification was based on absence of birefringence when the sample was viewed between crossed polarizers on a LM. Identification using TEM was based on shape of holes left in resin sections, the crystals themselves having either been dissolved in the preparation procedure or having fallen out during sectioning.
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RESULTS |
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Silica
Silica bodies are entirely absent from leaves of Philydraceae and Pontederiaceae (Prychid et al., 2003), and present in only some genera of Commelinaceae and Haemodoraceae (Table 1). Within Haemodoraceae, 20 species showed positive signs of silica accumulation and another three showed possible signs. All the species that contain silica are placed in the subfamily Conostylidoideae (Simpson, 1990, 1998); all genera of this subfamily are represented here. On the other hand, no silica was observed in Anigozanthos humilis (Conostylidoideae), or in any species of subfamily Haemodoroideae.
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Within Conostylidoideae, silica is present in the form of silica sand; i.e. numerous minute grains of silica. Individual grains vary in size, and sometimes coalesce into larger particles (Fig. 1A and B). Often the particles are present in drifts along one of the cell walls, perhaps due to a gravitational effect, or possibly an artefact of histological preparation. The distribution of silica deposition within the leaf varies between genera. In the distal (unifacial) part of the leaf (the leaf blade), in one genus (Anigozanthos) silica occurs entirely in the epidermal cells of the leaf blade and, in Blancoa and Macropidia, silica is present only in the bundle sheath cells in the leaf blade. However, silica is often more widespread in the proximal (bifacial) part of the leaf (the leaf sheath); e.g. in Blancoa and Conostylis, silica is also present in the adaxial epidermis in this region, although generally the greatest amount of deposition is found in the parenchymatous cells surrounding the vascular bundle (the bundle sheath). In some species (Blancoa canescens, Conostylis bracteata, C. setigera) all cells of the bundle sheath contained large amounts of silica, although rarely filling the cell entirely. Silica is apparently absent from lignified tissues, except in Conostylis setosa, in which the outer periclinal walls of the epidermal cells were lignified. However, it is possible that heavy staining of lignified tissues may mask the presence of silica sand.
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Anigozanthos.
Silica was present in all species except Anigozanthos humilis. In the leaf blade, silica sand is present only in epidermal cells, and occurs in small quantities, but in the majority of both adaxial and abaxial epidermal cells in the bifacial leaf sheath (Fig. 2A).
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Blancoa.
In the leaf blade silica is mostly absent from the epidermis, but present in the parenchymatous cells surrounding the vascular bundle (Fig. 2B); however, in the leaf sheath silica sand also occurs in smaller quantities in the adaxial epidermal cells, although it is apparently absent from the abaxial epidermal cells, which have thick lignified walls.
Conostylis.
The majority of species (except C. setigera, C. setosa and C. stylidioides) have fibre caps at the phloem poles of the vascular bundles, and (in all species except C. stylidioides) a lignified unifacial epidermis. Silica sand does not occur in the lignified tissue but is located primarily in the parenchymatous outer bundle sheath cells (Fig. 2C). In C. bealiana, silicified grains of silica, finer than those found in the bundle sheath cells, occur in the palisade mesophyll (Fig. 2D). Conostylis bracteata, C. setigera and C. setosa also have silica sand in palisade mesophyll cells. Small amounts occur in the spongy parenchyma in C. bealiana, C. bracteata, C. bromelioides, C. filifolia, C. juncea, C. setigera and C. setosa. The leaf sheaths of C. setosa contained sand in some unlignified adaxial epidermis cells. Occasionally a lignified adaxial epidermal cell was seen with a greater lumen diameter than those of the abaxial epidermis and silica grains were seen inside. Occasionally in Conostylis bealiana the silica sand appeared to have formed an almost spherical body of coalescent smaller units (Fig. 2E).
Macropidia.
Silica is rare but when present occurs in vascular bundle sheath cells in both the leaf sheath and leaf blade.
Phlebocarya.
Relatively large quantities of silica are present in both the parenchymatous vascular bundle sheath cells and the majority of epidermal cells. The epidermal cells are possibly lignified, the outer cell wall, possibly bearing a cuticle, appearing thicker than the inner. Small amounts also occur in the central mesophyll cells in P. filifolia and P. pilosissima and in subepidermal layers in P. filifolia. In the vascular bundle sheath cells of P. ciliata and P. filifolia, silica sand also occasionally coalesces into a spherical silica body (Fig. 2F). Silica was also seen in some of the sclerenchymatous bundle sheath cells in P. ciliata.
Tribonanthes.
Silica sand is rare but when present occurs mainly in the bundle sheath cells. In T. longipetala silica sand was seen occasionally in the lacunae and in the mesophyll cells.
Tapetal raphides
The distribution of crystals in vegetative tissues in Commelinales was reviewed by Prychid and Rudall (1999); raphides are generally present in vegetative tissues in these families. Crystal data on individual genera of the Haemodoraceae, however, is sparse, although raphides are known to occur in the family (Schulze, 1893; Dahlgren and Clifford, 1982). New data are presented here specifically on raphides that are present in the tapetum (Table 1). Tapetal raphides were observed here in some Haemodoraceae (A. flavidus, A. (A. bicolor x A. humilis) x A. flavidus, Conostylis candicans x aculeata) and some Comme linaceae (Tradescantia ohiensis, T. spathacea).
Haemodoraceae.
In Anigozanthos and Conostylis, raphide bundles occur in the tapetum surrounding the microsporocytes, dyads and early tetrads. As with Cochliostema odoratissimum (Hardy and Stevenson, 2000) raphides form in each tapetal cell prior to endothecium maturation and the onset of meiosis of the microsporocytes (Fig. 3A–D). The tapetal cells, at the microsporocyte stage, have a dense, granular protoplasm with numerous mitochondria. The crystals seem to form within vacuoles, with possibly one or two crystals per vacuole, within the protoplasm (Fig. 4A). The vacuoles can be dispersed throughout the cell and do not seem to be linked to each other so that the traditional tightly packed raphide bundle is not formed; however, all the crystals within a cell seem to follow the same orientation with respect to the axes of the cells. The bundles look like a collection of holes in transverse section, the crystals having fallen out, some holes are angular, and the crystals are long and needle-like in longitudinal section. SEM images show them to be blunt or wedge shaped at either end (Fig. 3D). The lengths of the tapetal raphides varies, with the maximum length being approx. 18 µm in Anigozanthos flavidus. No other crystal types are formed.
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At the dyad or early tetrad stage, numerous crystals, seemingly larger than those of the microsporocyte stage, occur per protoplast. Most protoplasts are still dense and grainy, but in some cases the protoplasmic material seems to be thinning out, although there are still numerous mitochondria present. During meiosis in the microsporocytes the cell walls of the tapetum are broken down. The protoplasts with their crystals fuse and then extend into the anther locule, forming a multinucleate plasmodium surrounding the free microspores until these mature (Fig. 4B and C). Groups of crystals often occur in vacuoles. Before anthesis the plasmodium degenerates. Tapetal material is deposited on the developing pollen grains forming the pollen coat.
Unlike that of Cochliostema odoratissimum (Hardy and Stevenson, 2000) the number of crystals remaining at anther dehiscence is large, suggesting that the crystals were not reabsorbed during pollen development.
Anthers had already dehisced in material examined of Tribonanthes longipetala, so raphides that were present in the anther locule could not conclusively be said to have originated from the tapetum.
Commelinaceae.
In Tradescantia ohiensis and T. spathacea, tapetal raphides are dispersed throughout the plasmodium surrounding the tetrads (Figs 4D and 5A) and again are associated with vacuoles. The raphides are more often single, not in bundles, and have a maximum length of approx. 14 µm in Tradescantia ohiensis. No other crystal types were seen.
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Philydraceae.
Anthers had already dehisced in material examined of Helmholtzia glaberrima, so raphides that were present in the anther locule could not conclusively be said to have originated from the tapetum.
Pontederiaceae.
In Monochoria africana, localized raphides occur at thin areas of the anther wall, possibly formerly the regions where the septum joined the anther wall to divide the anther into locules. The raphides project into the anther locule (Fig. 5B). Raphides are present in the layer adjacent to the tapetum, and thus could be derived from the tapetum, but this is unlikely. Occasionally styloids were also present in the anther locule, but these were very rare. No tapetal raphides were observed in Eichhornia azurea, Pontederia cordata var. cordata or P. lanceolata.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Presence or absence of characteristic cell inclusions may have systematic potential in commelinid monocotyledons, although the existing topology indicates de novo gains and losses in individual families (Fig. 6).
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Silica
This paper represents the first record of silica in Haemodoraceae. Thus, within Commelinales, it has been demonstrated that silica is present in leaves of two out of the four families (Haemodoraceae and Commelinaceae), but entirely absent from the other two families (Pontederiaceae and Philydraceae). In some phylogenetic analyses of commelinid monocots (e.g. Chase et al., 2000) the putatively basal family is Arecaceae (palms), in which silica bodies are invariably present (Tomlinson, 1961), often as rugose spherical bodies located in the bundle sheath cells (Prychid et al., 2003). Presence of silica bodies is putatively plesiomorphic for the ‘ZHC clade’, which consists of Zingiberales, Commelinales and Hanguana (Fig. 6). Silica bodies in Zingiberales are mostly more or less spherical with a rugose surface, as in palms, or occasionally conical, or ‘trough-shaped’ (i.e. rectangular with a central shallow depression) or present as silica sand. Similarly, as in Arecaceae, silica bodies in many Zingiberales occur in cells adjacent to the fibrous bundle sheath cells rather than the epidermis (Tomlinson, 1969), although they are also present in the epidermis in a few species (Prychid et al., 2003). Thus, the plesiomorphic conditions for silica bodies in the ZHC clade are probably spherical bodies located in the bundle sheath cells.
Within Haemodoraceae, silica is restricted to subfamily Conostylidoideae (sensu Simpson, 1990, 1998), almost always as silica sand, either in the epidermis or bundle sheath cells, or both. However, in Phlebocarya bundle sheath sand occasionally appears to form an aggregate spherical silica body. The occurrence of silica in Phlebocarya supports Simpson’s transfer of this genus from the Haemodoroideae to the Conostylidoideae Simpson (1990). Generally, the greatest concentration of silica in Haemodoraceae occurs in the vascular bundle sheath cells, Thus, silica of Haemodoraceae resembles that of Hanguana (Hanguanaceae) in both tissue location and granular morphology, although the two families are apparently not sister taxa (e.g. Graham et al., 2003). In Hanguana, silica bodies occur as irregular granular deposits, mainly in or near the vascular bundle sheath cells rather than in the epidermis (Solereder and Meyer, 1929; Smithson, 1956; Tomlinson, 1969).
The absence of silica bodies from Pontederiaceae and Philydraceae may indicate two separate losses of this character (Fig. 6), if this topology proves correct. Within Commelinaceae, silica bodies were earlier thought to be present only in the tribe Tradescantieae of the subfamily Commelinoideae (Tomlinson, 1969) but large silica infillings of the cell have now also been observed in Dictyospermum, of the tribe Commelineae (R. Faden, pers. comm., 2003). Silica bodies of the Tradescantieae are mainly small, spherical and spinulose and restricted to the epidermal cells (several per cell) (Molisch, 1918; Tomlinson, 1966, 1969), although silica sand has been observed in Zebrina pendula (Prychid et al., 2003).
Tapetal raphides
Tapetal raphides represent a rare character, only reported in a few monocot families of the order Commelinales (Table 1). Tapetal raphides were observed here in Anigozanthus and Conostylis (both Haemodoraceae), and Tradescantia (Commelinaceae), thus supplementing two earlier records in Haemodoraceae, in Lachnanthes (Simpson, 1988) and Xiphidium (Simpson, 1989), and in Philydraceae (Philydrum; Hamann, 1966) and Commelinaceae (Hardy and Stevenson, 2000). Their presence in the majority of families of Commelinales may therefore indicate an underlying cell chemistry that is preadaptive for their formation, perhaps related to regulation of free calcium levels in the tapetal cytoplasm.
The tapetum is a specialized layer of cells lining the anther locule and its development is linked with that of the adjacent sporogenous tissue. In monocots it is formed from periclinal division of the microsporangial secondary parietal layer. The tapetum functions as a nutritive tissue for the developing pollen grains, and produces exine precursors, orbicules, and sporophytic recognition proteins and lipids which form the pollen coat (Echlin, 1971; Esau, 1977; Pacini et al., 1985). In Haemodoraceae, Commelinaceae and some Pontederiaceae the tapetum is of the plasmodial (or amoeboid) type (Furness and Rudall, 1998). During meiosis in the microsporocytes the walls of the tapetum begin to break down. The tapetal protoplasts fuse and form a multinucleate plasmodium which enters the anther locule, and surrounds the tetrads and developing microspores. The plasmodium degenerates before the pollen grains are mature. The exact timing of plasmodium formation and degeneration may vary between taxa (Furness and Rudall, 1998).
Hamann (1966) found that in Philydrum (which has a secretory tapetum) almost every tapetal cell contained a raphide bundle; upon degeneration of the tapetum these are released between the developing pollen grains. Hardy and Stevenson (2000) also found that in Cochliostema odoratissimum (Commelinaceae) raphides formed in each tapetal cell prior to the onset of meiosis of the microsporocytes. Significantly lower quantities of prismatic or styloid-like crystals also occurred. Once the tapetal cell walls had broken down, the protoplasts and their crystals were found among the developing pollen grains. As the grains matured crystal quantities decreased and it was found that the remaining crystals had diminished in size. Hardy and Stevenson (2000) thus postulated that tapetal crystals had been reabsorbed during pollen development, thereby releasing free calcium possibly required in the ontogenetic process.
In mature anthers the crystals may easily be overlooked if the partitions between the anther locules have broken down so that anther wall raphides, or those of the ‘calcium oxalate package’ usually located at the anther stomium beneath the epidermis (D’Arcy et al., 1996), are also present in the anther locules. However, this would not apply in all cases; e.g. raphides only occur in the tapetal cells of Cochliostema odoratissimum and not in the surrounding wall layers (Hardy and Stevenson, 2000).
It has been shown that in some taxa (e.g. Gasteria; Tirlapur and Willemse, 1992; Willemse, 1993), during particular stages of microsporogenesis and pollen development there are significant increases in free calcium and activated calmodulin levels in the tapetum. However, apart from Commelinaceae, Haemodoraceae and Philydraceae, crystals are not formed in these cases. It would be interesting to compare the levels of free calcium just prior to crystal formation in tapetal cells of Commelinaceae, Haemodoraceae and Philydraceae, with those of Gasteria to test whether the levels are significantly higher in the crystal forming taxa. If the levels were significantly higher, this could explain the induction of crystal formation as a mechanism for regulation of cytoplasmic free calcium levels. Franceschi (1989) found that entire raphide bundles could be formed within 30 min of an increased calcium stimulus in Lemna minor. A similar process took 21–25 h in Yucca (Agavaceae) roots (Kausch and Horner, 1984).
Calcium is an essential substance for germinating pollen (Kwack and Brewbaker, 1961; Brewbaker and Kwack, 1963, 1964). Optimum pollen tube elongation also requires calcium ions. If the crystals adhere to the pollen grains when the anthers dehisced they could be transported along with the pollen to an appropriate stigma, where the crystals could dissolve in stigma secretion and help with pollen germination, although this requires further research.
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ACKNOWLEDGEMENTS |
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We are grateful to Robert Faden (Smithsonian Instiution, Washington) and Michael Simpson (San Diego State University) for critically reviewing the manuscript.
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