Molecular Plant Advance Access originally published online on July 17, 2009
Molecular Plant 2009 2(5):966-976; doi:10.1093/mp/ssp050
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Arabinan Metabolism during Seed Development and Germination in Arabidopsis
CNAP, Department of Biology, University of York, Heslington, York YO10 5YW, United Kingdom
1 To whom correspondence should be addressed. E-mail smm1{at}york.ac.uk, fax + 44 (0) 1904 328786, tel. + 44 (0) 1904 328775
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Abstract |
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 TOP Abstract INTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Arabinans are found in the pectic network of many cell walls, where, along with galactan, they are present as side chains of Rhamnogalacturonan l. Whilst arabinans have been reported to be abundant polymers in the cell walls of seeds from a range of plant species, their proposed role as a storage reserve has not been thoroughly investigated. In the cell walls of Arabidopsis seeds, arabinose accounts for approximately 40% of the monosaccharide composition of non-cellulosic polysaccharides of embryos. Arabinose levels decline to
15% during seedling establishment, indicating that cell wall arabinans may be mobilized during germination. Immunolocalization of arabinan in embryos, seeds, and seedlings reveals that arabinans accumulate in developing and mature embryos, but disappear during germination and seedling establishment. Experiments using 14C-arabinose show that it is readily incorporated and metabolized in growing seedlings, indicating an active catabolic pathway for this sugar. We found that depleting arabinans in seeds using a fungal arabinanase causes delayed seedling growth, lending support to the hypothesis that these polymers may help fuel early seedling growth.
Key Words: Cell walls • development • embryogenesis and seed development • Arabidopsis • arabinan • arabinose carbohydrate metabolism • physiology of plant growth
Received for publication May 1, 2009. Accepted for publication June 16, 2009.
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INTRODUCTION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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The plant cell wall provides mechanical support and serves as a point of communication and defence to plant cells, and, as such, it is essential in many aspects of growth and development. The primary cell wall provides a physical counterbalance to the turgor pressure in plant cells and also serves as the site at which rates and directions of plant cell expansion are controlled (Cosgrove, 2005). The cell wall is also the site of cell-to-cell adhesion and interaction in plants, and the apoplast is the medium for the transmission and perception of many signaling compounds (Somerville et al., 2004). The cell wall is a composite material incorporating a complex range of polymers, predominantly polysaccharides. These various polysaccharides appear to fulfill a range of structural functions. Cellulose microfibrils form the major structural elements of the wall, providing it with its inherent strength. These microfibrils are coated with matrix polysaccharides that fulfill functions as both spacing agents (keeping microfibrils apart) and linking elements (tethering microfibrils to one another), and this network serves as the primary site of action of proteins that modify cell wall extensibility during growth (Cosgrove, 2005). Pectic polysaccharides form a network coextensive with the cellulose/hemicellulose mesh and may be covalently linked to it (Popper and Fry, 2008). Typically, pectins are charged polysaccharides that are important in cell-to-cell adhesion, and also in cell wall elasticity (Jones et al., 2003).
Recent progress in cell wall research reveals that its function involves dynamic and complex regulatory mechanisms (Reiter, 2008). Since much of the carbon fixed in photosynthesis is incorporated into the cell wall, the changes in cell wall components should be, at least partially, affected by the fluxes of sugars occurring in the protoplast (Seifert, 2004). It has been shown that wall polysaccharides can serve as major carbohydrate storage reserves in the seeds of a number of plant species (Buckeridge et al., 2005; de Silva et al., 1993).
The structural complexity of the plant cell wall and the fact that its composition varies between different plant tissues and developmental stages reflect its varied functions in plant life (Knox, 2008). At the present time, we have developed a reasonable understanding of the range of polymers that are found in cell walls and of some of the interactions between these polymers (Somerville et al., 2004). However, our knowledge of the functional roles of particular wall polymers remains slight, and this is especially true of the pectins. Classically, pectins are characterized by the presence of galacturonic acid residues, and generally consist of homogalacturonan and rhamnogalacturonan I and II (RG-I, RG-II), both of which have complex branching structures (Mohnen, 2008). Arabinans are found in many cell walls and are generally considered to be part of the pectic network due to their extractability and the fact that they are often associated with RG-I, where, along with arabinogalactans, they form extensive side chains (Willats et al., 2001).
Whilst RG-I side chains account for much of the arabinan in plant cell walls, there are a number of instances in which arabinans appear to exist as independent polymers. These occurrences range from the apparently exotic, such as the cell walls of prickly pear spines, which are reportedly composed almost exclusively of equal amounts of cellulose and arabinan (Vignon et al., 2004), to the more familiar; for example, branched arabinans comprise the major non-cellulosic polymers of the cotyledons of some brassicas (Qouta et al., 1991; Rees and Richardson, 1966). These polymers differ significantly in their extractability from the cell wall in comparison to the side chains of RG-I. The complexity of cell wall matrix polysaccharides such as arabinans suggests a role for many enzymes in both their synthesis and degradation. Several enzymes capable of hydrolyzing arabinose-containing polymers have been described in microorganisms (Proctor et al., 2005), but only a few plant enzymes with this activity have been identified and their substrate specificity remains unclear. In particular, several plant 
-L-arabinofuranosidases belonging to glycosyl hydrolase families 3 and 51 (Fulton and Cobbett, 2003; Lee et al., 2003; Minic et al., 2006) have been partially characterized but their biological significance remains elusive. Interestingly, a recent characterization of the role of an -arabinofuranosidase (ARAF1) revealed pectic arabinans to be in vivo substrates of this enzyme (Chavez Montes et al., 2008).
In contrast, many biological functions have been proposed for arabinogalactan proteins (AGPs). Typically, AGPs incorporate a type II arabinogalactan. AGPs are implicated in many processes, including cell development, cell death, and cell-to-cell signaling, but the specifics of their mode of action remain elusive (Showalter, 2001).
More recently, soluble heteroglycans (SHGs), present in the cytosol of Arabidopsis leaves, have been identified as being rich in arabinose. It is proposed that these SHGs may be involved in the metabolism of plastid derived mono- and disaccharides in the cytosol (Fettke et al., 2005).
Whilst the structural relationships of arabinans in the cell wall are becoming clearer, there remains little understanding of their functional role. Studies using the LM6 antibody, which recognizes short linear stretches of (1–5), -L-arabinan, have revealed a range of associations of this epitope with, for example, meristematic cells but not elongating cells in carrot roots (Willats et al., 1999a, 1998) and tomato fruit cell walls prior to the onset of softening (Orfila et al., 2002). Other work has indicated a relationship between the presence of arabinans and cell-to-cell adhesion (Iwai et al., 2001; Orfila et al., 2001). Expression of a fungal arabinanase in transgenic potato plants led to a range of growth abnormalities including stunting, alterations in growth form, and infertility, suggesting that wall arabinans, or oligosaccharide signals generated from them, may have far-reaching developmental implications (Skjot et al., 2002). Other recent observations indicate important roles for plant arabinans: Jones et al. (2003) showed that cell wall arabinan is essential for the correct functioning of stomatal guard cell walls and proposed a role for arabinan in the maintenance of fluidity in the pectic network.
In this present work, we analyzed the role of arabinans during seed development and germination in Arabidopsis. Quantitative and qualitative studies revealed substantial changes in the abundance and distribution of this polysaccharide during seed establishment, and indicate that arabinose accumulates in a pronounced manner during seed development, and is then lost rapidly from the cell walls during germination and seedling establishment, when it may be metabolized by plant cells.
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RESULTS |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Arabinose Accumulates in Embryo Cell Walls and Declines Markedly during Germination
The mobilization of seed reserves fuels the morphological changes that will conclude with the establishment of an independent, autotrophic seedling. Beyond its clear involvement in the determination of cell shape and growth during post-germinative development, the cell wall plays a pivotal role in the carbon fluxes within the cell and, in some species, represents the major site of reserve deposition. In both mustard (Brassica napus) and cabbage (Brassica oleracea) arabinans are abundant in cotyledon cell walls of dry seed and are hydrolyzed during germination (Qouta et al., 1991; Rees and Richardson, 1966). Although the main metabolic reserves in Arabidopsis seeds comprise both fatty acids (in the form of triacylglycerols) and storage proteins, sugars also play an essential role in germination (Germain et al., 2001).
Examination of the monosaccharide composition of non-cellulosic cell wall polymers revealed that arabinose is the most abundant monosaccharide in embryos from mature seeds (Figure 1A) and its levels are notably higher than that seen in other tissues of Arabidopsis plants, such as leaves (Figure 1B). In embryo cell walls, arabinose is the most abundant monosaccharide from the non-cellulosic polysaccharides, accounting for approximately 40% of the total monosaccharide content compared with only 15% in rosette leaves (Figure 1A). Furthermore, an estimation of the arabinose content in the dry seed suggests that arabinose represents as much as 10–15% of the dry weight of Arabidopsis embryos.
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The arabinose content of cell walls showed marked changes during seed development and germination (Figure 1C). At torpedo stage (the earliest stage of embryo development that was measured), arabinose content was 20% of non-cellulosic sugars, and this increased steadily, reaching over 45% in dry seeds. Conversely, the arabinose content declined rapidly during germination to approximately 25% at 4 d after germination and 15% in mature leaves (Figure 1C).
Immunolocalization Studies Highlight Extensive Arabinan Remodeling during Seed Maturation and Germination
In dicots such as Arabidopsis, arabinose is found in a number of cell wall polymers, including RG-I, RG-II, and in the glycans of arabinogalactan proteins (Minic et al., 2006). The monoclonal antibody LM6 is specific in its recognition of at least five (1
5)--L-linked arabinose residues (Willats et al., 1998). Historically, these were found as side chains of RG-l, but, more recently, (15)--L-linked arabinan has been identified in cytosolic SHGs (Fettke et al., 2005). It was recently reported that LM6 recognizes epitopes in AGPs from the moss Physcomitrella patens (Lee et al., 2005), but, so far, there have been no reports of substantial linear (15)--L-arabinans in AGPs from higher plants. Other data indicate that most AGPs do not carry the LM6 epitope (Willats et al., 1998). Immunolabeling of Arabidopsis with LM6 antibodies was performed to examine arabinan distribution during seed development and germination. Immunofluorescent labeling of Arabidopsis embryos revealed a distinct trend in the distribution of the LM6 arabinan epitope, throughout seed maturation and germination (Figures 2, 3, and 4). Whilst the distribution of arabinan epitope varied between developmental stages, the labeling at each stage was consistent throughout all embryo tissues (Figure 3A). During the early stages of embryo development, walking stick, and bent cotyledon, LM6 labeling revealed a punctuate distribution of arabinans within the cytoplasm of all cells and, to a lesser extent, at the cell wall (Figure 2A and 2C). A few larger areas of cytoplasmic fluorescence were also observed. As seeds reached maturity, LM6 labeling at the cell wall was more intense, but patchy (Figures 2E and 3B). The intensity of LM6 labeling was compared across developmental stages by the preparation of all material and immunolabeling in parallel, and by comparative imaging using the same confocal configuration (not shown). However, images presented here have been obtained at the optimal confocal configuration (optimal laser power and detector gain) for each sample and were taken in parallel with a negative control (no primary antibody—data not shown) at the same configuration to ensure that labeling was specific.
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In addition to wall labeling, the intracellular labeling seen with LM6 was at its most intense in mature seed (Figure 3B). However, the mature seed also showed a considerable amount of autofluorescence. To distinguish antibody labeling from autofluorescence, samples were spectrally unmixed (where the fluorescent profile of autofluorescence is distinguished from that of the fluorescent antibody conjugate, in this case, FITC). This confirmed that the LM6 label was found in a patchy distribution at the cell wall, and in larger fluorescent regions in the cytoplasm. The background autofluorescence indicates that LM6 appeared to label parts of large structures in the cytoplasm. In order to identify a more exact sub-cellular location of the LM6 epitope at this stage, ultra thin sections of mature embryo were immunolabeled with LM6 and 10 nm gold and viewed with an electron microscope (Figure 3C). Consistent with immunofluorescence images, LM6 labeling was concentrated at the cell periphery, particularly at the inner cell wall (Figure 3C, cw) at regions of apparent vesiculation (3C, v). However, indirect immunogold labeling failed to identify the more central locations in the cytoplasm, where LM6 labeling was observed with immunofluorescence (Figure 3B). This was despite optimizing the labeling procedure for immunogold, and confirming the specificity of LM6 labeling at the cell periphery by way of a negative control (Figure 3D). No unusual structures were revealed by these ultrastructural studies, which highlighted an accumulation of electron transparent structures, most likely to be lipid bodies, at this stage (Figure 3D, *).
At 24 h post germination, LM6 labeling in the cytoplasm was considerably reduced (Figure 4A), and very few of the large areas of cytoplasmic fluorescence, characteristic of mature embryos, were observed. Conversely, LM6 labeling at the cell wall was increased, and appeared less patchy than that observed in mature embryos (Figure 3B). Rather, the arabinan epitope appeared to be present consistently along all lengths of the plasmalemma face of the cell wall. By 48 h post germination, a dramatic reduction in LM6 labeling was apparent, and the arabinan epitope was detected only at the external surface of distal root cap cells (Figure 4C). In root tissues, away from the root cap, cell wall-localized LM6 labeling was most abundant in vascular tissue (Figure 4C). This pattern of arabinan distribution remained unchanged at 72 h after germination (Figure 4E).
Immunolabeling of galactan and xyloglucan (not shown) and partially methyl-esterified homogalacturonan with JIM7 (Knox et al., 1990) was undertaken to determine whether the pattern of labeling seen with LM6 was representative of cell wall polysaccharides or specific to arabinan. JIM7 labeling was located specifically at the cell wall at all three stages of embryo development (Figure 2B, 2D, and 2F), but was restricted to the vascular tissue in mature embryos (Figure 2F). Negative controls (not shown) revealed that the cytoplasmic fluorescence in mature embryos was autofluorescence, not JIM7 labeling. Some specific punctuate staining was observed at the walking stick and bent cotyledon stages with JIM7 (Figure 2B and 2D), but to a lesser extent than that seen with LM6 (Figure 2A and 2C). At 24 h after germination, JIM7 labeling was found predominantly at the cell wall of root caps cells and vascular tissue (Figure 4B). As with mature seed, negative controls revealed cytoplasmic labeling to be non-specific autofluorescence. In contrast to LM6 labeling, which declines 48 h after germination (Figure 4C and 4E), the homogalacturonan epitope labeled by JIM7 was most abundant at 48 and 72 h after germination, when it was detected at the cell wall of all root cells (Figure 4D and 4F).
Arabinanase Treatment of Embryos Reduces Seedling Growth
In order to determine whether reducing arabinan content in embryos prior to germination had any effect on germination and growth, Arabidopsis embryos were removed from seed coats and exposed to exogenous arabinanase and arabinofuranosidase treatment, before germination (Figure 5). Treatment of embryos with either endoarabinanase or arabinofuranosidase both resulted in retarded growth of the seedlings (Figure 5A). The effect of treatment of seed with A. niger endoarabinanase was particularly strong, perhaps underlining the importance of arabinans rather than single arabinosyl residues in the effects observed. In all cases, the experimental conditions were optimized to ensure that the effect on the germination rate of the seeds was negligible (see Methods and Figure 5B).
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Arabinose Is Metabolized in Arabidopsis Seedlings and Enters Mainstream Metabolic Pathways
The dramatic changes in arabinan content and localization after germination indicate the presence of metabolic machinery dedicated to the synthesis and turnover of these polysaccharides, but, currently, little is known about the enzymes involved in the synthesis and degradation of arabinans. An arabinosyl transferase gene (ARAD1) has been characterized in Arabidopsis, and plants carrying a knockout insertion in this gene exhibit only 30% of arabinose in RG-I (Harholt et al., 2006). While no endoarabinanases have been described in plants, enzymes with arabinofuranosidase activity have been described recently (Minic et al., 2004, 2006), and mutants with altered arabinofuranosidase expression show cell wall modifications and impaired germination (Chavez Montes et al., 2008). In order to establish whether Arabidopsis seedlings can metabolize arabinose as a carbon source during germination, a series of feeding experiments were performed using 14C-labeled arabinose in order to monitor its metabolic fate. The level of 14CO2 release increased linearly with time for several hours after the seedlings were fed with labeled arabinose (Figure 6A). This observation demonstrates the ability of seedlings to use arabinose in metabolism and respiration. After 12 h of incubation, approximately 30% of the arabinose was metabolized and incorporated in a range of different compounds (Figure 6B). Whilst almost 50% of the consumed 14C was released as CO2, significant quantities were also incorporated into various cellular components and metabolic intermediates. After 12 h of incubation with 14C arabinose, 29% was associated with protein, 13% with cell walls, 3% with starch, and 7% with soluble metabolites. These data indicate that free arabinose is readily taken up by seedlings and enters main stream metabolism, making it plausible that arabinose mobilized from the wall is used as a metabolic sugar by the plants.
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DISCUSSION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Arabinans Are Abundant in the Cell Walls of Arabidopsis Embryos and Undergo Extensive Degradation during Germination
The large changes observed in number and localization of arabinans in Arabidopsis seeds suggest that these cell wall components could play a role linked to the establishment of germinating seedlings. Interestingly, arabinans have been reported to be abundant polymers in the cell walls of seeds from a range of plant species (Dourado et al., 2004; Eriksson et al., 1996; Navarro et al., 2002), but their potential role during germination has been largely ignored. Studies amongst the brassicas have demonstrated the abundance and subsequent breakdown of these polymers during germination in both cabbage and mustard (Qouta et al., 1991; Rees and Richardson, 1966). Our studies of the monosaccharide composition of non-cellulosic polysaccharides of the cell walls of Arabidopsis embryos in seeds and during germination indicate that arabinose accounts for approximately 40% of the monosaccharide composition of non-cellulosic polysaccharides of embryo cell walls, and these levels decline to approximately 15% during seedling establishment.
There is even evidence for substantial levels of pectic arabinan in the embryos of cereals, which typically have very low levels of pectin in their walls. For example, Gibeaut et al. (2005) showed that at the start of germination, barley coleoptile cell walls have similarly high levels of pectic arabinose to those we report here in Arabidopsis, and that these decline sharply during germination.
Since the second most abundant monosaccharide in seeds is xylose, the possibility that other arabinose-rich polysaccharides such as arabinoxylan could follow the same dynamic suggested here for arabinans should be considered. However, unlike arabinose, xylose levels remain fairly constant, even at 15 d after flowering (data not shown).
Our immunolocalization experiments show that linear arabinans accumulate during embryo development and decrease 48 h after germination, and that this includes a loss of material from the cytoplasm as well as the cell wall. Whilst this correlates well with the arabinose trend shown by monosaccharide analysis, it is worth qualifying that the LM6 epitope specifically recognizes 1–5 arabinan, which is only known to be associated with RG-l and SHGs in dicots (Fettke et al., 2005; Willats et al., 2001). At the early stages of embryo development, the cytoplasmic distribution of LM6 epitopes is manifest as small punctuate regions with a distribution similar to that seen in Arabidopsis and other root material labeled with JIM84, a monoclonal antibody that recognizes an epitope located at the Golgi (Jin et al., 2001). No co-localization experiments were undertaken here, but it seems likely that the labeling we see at these early stages may be Golgi-localized.
In mature embryos, the cytoplasmic labeling is evident as larger fluorescent regions. Whilst indirect immunogold labeling did not disclose the location of this epitope, it may be present in soluble heteroglycans, which have recently been reported to be rich in arabinose with 1–5 linkages (Fettke et al., 2005). It is curious that the large cytoplasmic labeling observed with LM6 antibodies by immunofluorescence were not detected by immunogold labeling with the same antibodies, and this is something that requires further studies. We propose that the localization of arabinan to vesiculated regions at the cell wall is perhaps consistent with the deposition of cell wall material, since the onset of germination is accompanied by a considerable increase in the LM6 epitope at the cell wall, and a marked reduction in cytoplasmic labeling. Whilst it is also possible that these two regions may represent distinct sets of macromolecules, both containing linear (1–5) arabinan, immunolabeling studies with JIM7, which recognizes a homogalacturonan epitope, indicate that the distribution seen with LM6 does not follow a trend seen with other pectic polysaccharides. Additionally, whilst immunolabeling of galactan and xyloglucan (with LM5 and LM15/CCRC-M1, respectively, results not shown) both showed an accumulation of labeling at the cell wall of mature embryos, the dynamics of distribution varied from that seen with arabinan, and neither showed the extensive cytoplasmic labeling seen with LM6 in mature embryos. By 48 h after germination, the cytoplasmic labeling seen with LM6 in mature embryos is no longer apparent, perhaps indicating that the polymer it is associated with is rapidly metabolized, whilst the loss of LM6 labeling in the cell wall takes longer to disappear. The persistence of the arabinan epitope at the external surface of distal root cap cells after 48 h correlates well with surface labeling of 15-d Arabidopsis roots where LM6 is observed at the distal root apex (Willats et al., 2000), and also 10-d carrot roots, where LM6 is found specifically at the root apex (Willats et al., 1999b).
Degradation of Seed Arabinan Leads to Delayed Seedling Growth
Cell wall composition varies greatly within a plant from tissue to tissue and between developmental stages. Since the cell wall composition is likely to reflect its functional roles at a given moment, it seems likely that the abundance of arabinans in embryo cell walls reflects a biological function for this polymer. For example, it has been described that the presence of arabinan in the pectin network is essential to confer the flexibility required by guard cells to perform the changes in size and shape necessary for stomatal opening (Jones et al., 2005). Similarly, it has been shown that enzymatic modification of pectic arabinans is required for normal mucilage release (Arsovski et al., 2009).
During development, the Arabidopsis embryo increases from a few hundred cells at the globular stage to approximately 20 000 cells at maturation. Similarly, the volume of the embryo increases approximately 20 times during the maturation stage (Gomez et al., 2006). After germination, it is necessary for the embryo to overcome the mechanical resistance of the seed teguments in order to rupture the testa and endosperm, and this involves substantial cell expansion. These processes require a considerable degree of cell wall plasticity and volume adjustment. It is possible that the arabinan deposited in the cell wall during embryo development might represent a conditioning that is necessary to permit the high degree of cell wall flexibility required during germination. This proposition is supported by the delay in growth observed after treatment of seed with fungal arabinanase.
Do Arabinans Serve as a Storage Reserve?
The mobilization of storage reserves during the early stages of seedling growth is a prerequisite for successful establishment prior to the initiation of photosynthesis. Lipids represent the major seed reserve in oilseed plants such as Arabidopsis, and mutants unable to mobilize storage lipids generally require sucrose supplementation for normal seedling establishment (Germain et al., 2001). Arabidopsis seeds do not contain starch (Hills, 2004) and plants with lesions in the glyoxylate cycle and gluconeogenesis are compromised in their ability to germinate (Eastmond et al., 2000). However, it is possible that there may be other sources of sugars in Arabidopsis seeds. In some legume species, cell wall polysaccharides such as galactomannans serve as major storage reserves (Reid, 1985), and there is evidence to suggest that a flux of sugars from the cell wall into central metabolism also happens under stress conditions. A well characterized group of genes induced under dark conditions includes several cell wall-associated glycosyl hydrolases (Fujiki et al., 2001). Sugar depletion, on the other hand, has been shown to induce the secretion of three cell wall hydrolases into the apoplast, leading to reduced monosaccharide content of the matrix polysaccharide fraction (Lee et al., 2004).
The 14CO2 release observed when germinating Arabidopsis seeds are incubated in the presence of radiolabeled arabinose indicates that these seedlings are able to use this sugar as a respiratory substrate (Figure 6). Furthermore, the radiolabeled carbon is incorporated into several major cellular components. Similar incorporation in ethanol soluble and insoluble fractions was obtained when 5-day-old seedlings were incubated with L-[3H] arabinose (Dolezal and Cobbett, 1991). An arabinose salvage pathway has been proposed, where arabinose released from the cell wall is reincorporated into polysaccharides via phosphorylation, but no information is available on how arabinose is interconverted and incorporated into the central metabolism. Arabinose metabolism has received little attention in plants, but has been studied in plant-degrading bacteria, where three enzymes are required to enable L-arabinose to enter the pentose phosphate pathway: L-arabinose isomerase (converts arabinose to ribose), L-ribulose kinase (produces ribulose 5-phosphate), and L-ribulose 5-phosphate epimerase, which produces d-xylulose 5-phosphate, a key intermediate in the pentose phosphate pathway (Sa-Nogueira et al., 1997). Although there are potential homologs of all three of these genes in the Arabidopsis genome (e.g. At4g30310 for the AraB ribulose kinase or At5g61410 for ribulose-5-phosphate epimerase kinase), insertion lines for these genes do not show any phenotype associated with sugar depletion (data not shown).
Interestingly, arabinans have been reported to be abundant polymers in cell walls of seeds from a range of plant species (Dourado et al., 2004; Eriksson et al., 1996; Navarro et al., 2002); however, their potential role as a storage reserve has been largely ignored. Early reports on the isolation of arabinan active enzymes in carrot cell cultures suggest the possibility of self-degradation of the pectic matrix in these cells (Konno et al., 1987), and the mobilization of arabinans from coleoptile cell walls in barley during the first 48 h of germination represents evidence of this process occurring in a grass species (Gibeaut et al., 2005).
These observations indicate that arabinose levels may be relatively high in seeds across a wide range of plants, suggesting a conserved function for arabinans in this context. The ability to survive desiccation and subsequent rehydration is a key aspect of seed function. In this context, it is interesting to note that the cell walls of resurrection plants (that can survive desiccation of their vegetative tissues) are notably rich in arabinose (Moore et al., 2009). Rees and Richardson (1966) suggested that arabinans might be important in maintaining flexibility in mustard seeds during desiccation and germination, and, more recently, Jones et al. (2003) showed that arabinans play an important role in the flexibility of stomatal guard cell walls. Thus, it is possible that arabinans may play a role in cell walls associated with seed desiccation and that after germination, they are metabolized because they are no longer required for this role, except perhaps in specialized tissues such as the root cap and meristem, where immunolocalization studies indicate them to be abundant (Willats et al., 2000, 1999b).
The large quantities of arabinans in Arabidopsis embryos and their subsequent metabolism during germination are highly suggestive of some specialized function, and it is clear that further work is required to probe the nature of this specialization.
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METHODS |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Monosaccharide Analyses in Embryos and Germinating Seeds
Cell wall monosaccharide analysis was performed in triplicate with samples composed of 100 embryos, seeds, or germinating seedlings. Immature Arabidopsis thaliana (Columbia) seeds were taken from siliques at 8, 12, and 15 d after flowering, the embryos removed from the teguments, and cell walls extracted as described below. Samples for mature seeds were prepared by removing the seed teguments after imbibing the seed for 24 h at 4°C. For germinating seed samples, the seeds were stratified for 4 d at 4°C, transferred to growth rooms at 20°C, and harvested after 24, 48, and 72 h. Seed teguments were removed and cell wall from seedlings prepared. Cell walls were prepared by homogenizing plant materials in liquid phenol and washing with chloroform:methanol (2:1) before sedimentation by centrifugation. The pellets were washed twice with 95% ethanol and left to dry. To analyze the monosaccharide content of non-cellulosic polysaccharides, wall material was hydrolyzed with 2 M TFA for 4 h at 100°C. Monosaccharide analyses were performed by HPAEC using a Dionex Carbopac PA-10 as described in Jones et al. (2003). The monosaccharide standards used for quantification were arabinose, fucose, galactose, galacturonic acid, glucose, glucuronic acid, mannose, rhamnose, and xylose.
Treatment of Seed with Fungal Arabinanase
Sterile Arabidopsis seeds were placed in microcentrifuge tubes and treated with 10 U of previously dialysed Aspergillus niger endoarabinanase or arabinofuranosidase (Megazyme) in water for 16 h at room temperature. Controls were processed in the same way; seed was maintained in water. After enzymatic treatment, the seeds were rinsed and dried on sterile filter paper for 72 h at room temperature. Once dried, the seed was sown on 
MS plates and germination was scored after 5 d. The experiments were done in quadruplicate using 50 seeds/sample.
Microscopy
Preparation of Arabidopsis embryos and germinating seed for immunocytochemistry
Arabidopsis (Col-0) embryos at torpedo, bent cotyledon, and mature (not dried) developmental stages were prepared for microscopy following the removal of the seed coat. For germinating seedlings, Arabidopsis seed was sterilized by chlorine vapor and plated onto 0.8% plant agar with half-strength MS Basal Salt Mixture, pH 5.7. Seed was stratified at 4°C for 4 d before growing for 24, 48, or 72 h at 24°C with a 16-h day.
Whole embryos/seedlings were fixed under vacuum for 1 h at ambient temperature in 0.1 M sodium cacodylate buffer, pH 7.0, containing 1% glutaraldehyde, 2% paraformaldehyde, 0.1 M calcium chloride, and 0.5% (w/v) sucrose. Samples were processed using a progressive low-temperature method (Vandenbosch, 1991) and embedded in LRWhite resin containing 0.5% benzoin methyl ether. Samples were embedded in gelatin capsules and the resin was polymerized with ultraviolet light (–20°C for 24 h followed by –10°C for 24 h).
Immunofluorescence confocal microscopy
Semi-thin (500 µM) sections of resin-embedded material were cut using a Leica Ultracut UCT and dried onto Vectabond coated slides. All reagents were filter sterilized before use. Sections were blocked with 3% bovine serum albumin (BSA) in phosphate buffered saline pH 7.4 (PBS) for 30 min before incubation with primary antibody (JIM7 or LM6 diluted 1:10 in 1% BSA/PBS) or negative control (1% BSA/PBS) for 1 h at ambient temperature. Sections were washed with PBS before incubation in secondary antibody (anti-rat IgG whole molecule: FITC conjugate diluted 1:40 with 1% BSA/PBS) for 1 h at ambient temperature in the dark. Sections were washed before mounting with Citifluor antifade (glycerol/PBS). Samples were viewed with a Zeiss LSM 510 Meta on an Axiovert 200. Fluorescence was excited with a 488 laser and emission was filtered using a 505–530 bandpass. For spectral unmixing, images were collected between 497 and 604 nm in 10.7-nm increments using the spectral metahead. Autofluorescence and FITC emission profiles, made using unlabeled samples and concentrated conjugated secondary antibody, respectively, were used to separate the collected data.
Antibodies
JIM7 (which recognizes partially methyl-esterified epitopes of homogalacturonan (Knox et al., 1990)), LM6 (which recognizes a linear pentasaccharide in (15)--L-arabinans, (Willats et al., 1998)), LM5 (which a recognizes a linear tetrasaccharide in (1–4)-b-D-galactan (Jones et al., 1997)), and LM15 (which recognizes the XXXG motif of xyloglucan in Arabidopsis (Marcus et al., 2008)) were gifts from Professor Paul Knox, University of Leeds, UK, and are available commercially from PlantProbes.
CCRC-M1 (which recognizes the fucose residue of xyloglucan (Puhlmann et al., 1994)) was obtained from the Complex Carbohydrate Research Center, University of Georgia, USA (LM5, LM15, CCRC-M1, results not shown).
Indirect immunogold labeling
Pale gold (70–90 nm) ultra-thin sections were cut with a Diatome diamond knife, using a Leica Ultracut UCT, and mounted on 200 hexagonal mesh nickel grids. For immunogold labeling, grids were inverted onto 30-µl droplets of filter-sterilized reagent, as follows: Blocker (3% BSA in PBS containing 0.1% Tween 20; PBST) for 30 min, primary antibody (LM6 diluted 1:10 in 1% BSA/PBST), or control (1% BSA/PBST) for 1 h at 30°C, wash (PBST), secondary antibody (anti-rat IgG:10nm Gold conjugate diluted 1:20 with 1% BSA/PBST) for 1 h at 30°C, wash (PBST, ultrapure water). Sections were post-stained with 2% aqueous uranyl acetate, then lead citrate (Reynolds, 1963) in a carbon dioxide-free chamber. Indirect immunogold labeling was viewed using a FEI Technai G2 TEM operating at 120 kV. Images were captured using AnalySIS software and a Megaview III CCD camera.
14C Arabinose Feeding
Four hundred mg of sterilized Arabidopsis seeds (ecotype Columbia) were stratified for 4 d at 4°C on MS media and allowed to germinate for 24 h in a 12-h light/dark cycle at 20°C. The germinating seeds were subsequently transferred to airtight vials containing a small tube with 200 µl 2 M KOH, in order to quantify the 14C released. Assays were started by adding 3 ml of 500 µM labeled arabinose (0.3 KBq ml–1). The resulting K214CO3 was placed in scintillation fluid and determined in liquid scintillation counter. After feeding with labeled arabinose, the seed was ground in ethanol and fractionated into the ethanol soluble fraction (soluble sugars) and ethanol insoluble matter. The ethanol insoluble fraction, in turn, was removed and sequentially digested with macerozyme (25 U ml–1, Hepes pH 7.0, 12 h), amyloglucosidase (30 U ml–1, in acetate buffer pH 4.8, 12 h), proteinase K (90 U ml–1 in Hepes pH 7.0, 12 h) to obtain the label incorporated into the starch and protein fractions. Finally, the cell wall material that remained insoluble after these treatments was re-suspended in scintillation liquid and determined in the liquid scintillation counter.
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We acknowledge the financial support of the BBSRC.
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We thank Professor Paul Knox, University of Leeds, UK, for the gift of LM5, LM6, LM15, and JIM7 antibodies. No conflict of interest declared.
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