Molecular Plant Advance Access originally published online on August 20, 2009
Molecular Plant 2009 2(5):943-965; doi:10.1093/mp/ssp061
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Xyloglucans of Monocotyledons Have Diverse Structures
School of Biological Sciences, The University of Auckland, Private Bag 92019, Auckland, New Zealand
1 To whom correspondence should be addressed. E-mail p.harris{at}auckland.ac.nz, fax +64-9-373-7417, tel. +64-9-373-7599, ext. 88366.
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Abstract |
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Except in the Poaceae, little is known about the structures of the xyloglucans in the primary walls of monocotyledons. Xyloglucan structures in a range of monocotyledon species were examined. Wall preparations were isolated, extracted with 6 M sodium hydroxide, and the extracts treated with a xyloglucan-specific endo-(1
4)-β-glucanase preparation. The oligosaccharides released were analyzed by high-performance anion-exchange chromatography and by matrix-assisted laser-desorption ionization time-of-flight mass spectrometry. Oligosaccharide profiles of the non-commelinid monocotyledons were similar to those of most eudicotyledons, indicating the xyloglucans were fucogalactoxyloglucans, with a XXXG a core motif and the fucosylated units XXFG and XLFG. An exception was Lemna minor (Araceae), which yielded no fucosylated oligosaccharides and had both XXXG and XXGn core motifs. Except for the Arecales (palms) and the Dasypogonaceae, which had fucogalactoxyloglucans, the xyloglucans of the commelinid monocotyledons were structurally different. The Zingiberales and Commelinales had xyloglucans with both XXGn and XXXG core motifs; small proportions of XXFG units, but no XLFG units, were present. In the Poales, the Poaceae had xyloglucans with a XXGn core motif and no fucosylated units. In the other Poales families, some had both XXXG and XXGn core motifs, others had only XXXG; XXFG units were present, but XLFG units were not.
Key Words: Commelinid monocotyledons • non-commelinid monocotyledons • plant cell walls • Poaceae • xyloglucans • xyloglucan-specific endo-(1 4)-β-glucanase
Received for publication April 29, 2009. Accepted for publication July 10, 2009.
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INTRODUCTION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Xyloglucans are polysaccharides that occur in the primary cell walls of all angiosperms (flowering plants), although the proportions vary (Fry, 1989; Hayashi, 1989; Harris, 2005). In eudicotyledons, they often comprise 20–25% of the dry weight of the walls, but, in some species, such as celery (Apium graveolens), they comprise only 2% (Fry, 1989; Thimm et al., 2002). Small proportions of xyloglucans, usually
2–5%, also occur in the walls of the monocotyledon family Poaceae (grasses and cereals) (Fry, 1989). At least some of the xyloglucans are hydrogen bonded to the cellulose microfibrils in the walls and may cross-link adjacent microfibrils, so constraining cell enlargement (Bootten et al., 2004). Xyloglucans are thus thought to play a role in controlling cell enlargement (Rose et al., 2002; Fry, 2004). However, in addition to primary walls, xyloglucans occur as the major component of thick, non-lignified secondary walls in the cotyledons or endosperms of seeds in some species of the family Fabaceae as well as some other eudicotyledon families, but not in monocotyledons (Kooiman, 1960; Harris, 2005). These seed xyloglucans function as reserve carbohydrates and are mobilized during germination (Buckeridge et al., 2000).
Structurally, xyloglucans are composed of a backbone of β-D-Glcp residues linked (1 4)-, with 
-D-Xylp residues attached at O-6 to a proportion of these Glcp residues. Other substituents are also present on some of the -D-Xylp residues. To help describe the structures of xyloglucans, Fry et al. (1993) developed an unambiguous nomenclature with the letters G, X, S, L, and F referring to the following structures: G = unsubstituted β-D-Glcp; X = -D-Xylp-(1 6)-β-D-Glcp; S and L = X with -L-Araf-(1 2)- and β-D-Galp-(1 2)- attached, respectively; and F = L with -L-Fucp-(1 2)- attached (Figure 1). A common approach to analyzing xyloglucans structurally is to treat them with either a cellulase (endo-(1 4)-β-glucanase), or more recently with a xyloglucan-specific endo-(1 4)-β-glucanase, and identify and quantify the oligosaccharides released (Vincken et al., 1997). The enzymes are unable to hydrolyze the backbone where a β-D-Glcp residue is substituted, so the sequence ...GXXXGX... yields the oligosaccharide XXXG (Hoffman et al., 2005).
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The xyloglucans in the thick, non-lignified walls of seeds have been investigated in a number of species and have all been found to have similar structures (Buckeridge et al., 1992, 2000; Harris, 2005). For example, treatment of the xyloglucans from the cotyledons of tamarind (Tamarindus indica, Fabaceae) seeds with cellulase released four types of oligosaccharides: XXXG, XLXG, XXLG, and XLLG (Buckeridge et al., 1992; Marry et al., 2003). Thus, these xyloglucans contain galactose and have a repeating XXXG core motif; they are referred to as galactoxyloglucans (Figure 1). The xyloglucans that occur in the primary walls of most eudicotyledons also have a repeating XXXG core motif, but contain fucose in addition to galactose and are referred to as fucogalactoxyloglucans (Figure 1). Treatment with cellulase or a xyloglucan-specific glucanase yields the three major oligosaccharides: XXXG, XXFG, and XLFG, with smaller proportions of XXLG, XLLG, and XLXG (Harris, 2005). The proportions of the different oligosaccharides vary somewhat with the organ from which the walls were isolated (Pauly et al., 2001).
However, fucogalactoxyloglucans are not the xyloglucans in the primary walls of all eudicotyledons. Most known structural variation in eudicotyledon xyloglucans occurs in a group of plants known as the asterids (APG II, 2003; Harris, 2005; Hoffman et al., 2005; Hilz et al., 2007). Within this group is the economically important family Solanaceae and the structures of the xyloglucans of species in this family have been especially well examined. These xyloglucans have S- but no F-containing subunits, and are often referred to as arabinoxyloglucans (Figure 1). Unlike the galactoxyloglucans and the fucogalactoxyloglucans, arabinoxyloglucans have a repeating XXGG core motif. For example, when treated with a xyloglucan-specific endo-(1 4)-β-glucanase, the xyloglucans of the primary walls of tobacco (Nicotiana tabacum) leaves yielded mostly XXGG and XSGG and only very small proportions of L-containing oligosaccharides (Hoffman et al., 2005).
Compared with the xyloglucans in the primary walls of eudicotyledons, much less is known about the structures of the xyloglucans in the primary walls of monocotyledons. Most work has been done on the structures of the xyloglucans in the family Poaceae (grasses and cereals). These xyloglucans contain XXGG, XXGGG, and XXGGGG as common repeating core motifs, with small proportions of XXG and XXGGGGG also present (Kato et al., 1981, 2004a; Fry, 1989; Gibeaut et al., 2005). We therefore refer to the repeating core motif as XXGn, where n = 1–5 (Figure 1). Galactose-containing oligosaccharides such as XLGGG and LXGGG are present in Poaceae xyloglucans (Kato et al., 2004a), but small amounts of fucose-containing oligosaccharides have been reported only once (McDougall and Fry, 1994).
The Poaceae form part of a large group of monocotyledons, the commelinids, first recognized by the presence of ester-linked ferulic acid in their primary walls (Harris and Hartley, 1980; Rudall and Caddick, 1994; Harris et al., 1997; Harris, 2005). Subsequent phylogenetic studies of monocotyledons using the nucleotide sequences of genes resolved the same group as a monophyletic clade (Chase et al., 1993, 2006). The commelinids comprise four orders: the Poales (grasses and cereals, rushes, sedges, bromeliads, etc.), Commelinales (spiderworts, etc.), Zingiberales (gingers, etc.) and Arecales (palms) (APG II, 2003). In addition, the presence of ester-linked ferulic acid in their primary cell walls resulted in the identification of the genera Baxteria, Calectasia, Dasypogon, and Kingsia as commelinid monocotyledons. As a result, the genera were reclassified as the family Dasypogonaceae, although the order in which they should be placed is still uncertain (Rudall and Caddick, 1994; Rudall and Chase, 1996). As now defined (APG II, 2003), Poales (sensu lato) is a large order of 18 families, but formerly (Dahlgren et al., 1985) Poales (sensu stricto) was a group of six related families, including the Poaceae, now referred to as the graminoid clade. Except for a report suggesting that pineapple (Ananas comosus) fruit xyloglucan is a fucogalactoxyloglucan, nothing is known about the structures of the xyloglucans of commelinid monocotyledons outside the Poaceae (Kato et al., 2001).
Other monocotyledons, which we refer to as non-commelinids, comprise the Acorales (sweet flag), Alismatales (aroids and various aquatic plants), and the liliid orders Asparagales (asparagus, onion, etc.), Liliales (lilies, etc.), Pandanales (screw-pines, etc.), Dioscoreales (yams), and Petrosaviales. Unlike the commelinids, the non-commelinids have no, or very small amounts of, ester-linked ferulic acid in their primary walls (Harris and Hartley, 1980; Harris et al., 1997; Harris, 2005). In contrast to Poaceae xyloglucans, those of onion (Allium cepa), garlic (A. sativa), and their hybrid (family Alliaceae) are fucogalactoxyloglucans, with structures similar to those in the primary walls of most eudicotyledons (Ohsumi and Hayashi, 1994).
Here, we report the results of a study of the structures of xyloglucans in the primary walls of a range of both commelinid and non-commelinid monocotyledons and discuss the results in a phylogenetic context. We used a xyloglucan-specific endo-(1 4)-β-glucanase preparation to release xyloglucan oligosaccharides, which were then analyzed. Instead of treating whole cell walls, an alkali extract of the walls was used to avoid potential problems of xyloglucan accessibility. Another advantage of using such an extract was that O-acetyl groups, which commonly occur on xyloglucans (Figure 1), were removed, so reducing the number of different oligosaccharides and simplifying their analysis. The xyloglucan oligosaccharides were analyzed using two rapid methods that give profiles (‘fingerprints’) of the oligosaccharides: high-performance anion-exchange chromatography (HPAEC) with pulsed amerometric detection (PAD) and matrix-assisted laser-desorption ionization time-of-flight mass spectrometry (MALDI–TOF MS). A similar approach has previously been used to rapidly structurally phenotype Arabidopsis thaliana wall mutants (Lerouxel et al., 2002).
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RESULTS |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Enzymatic Activity of the Xyloglucan-Specific Endo-(1
4)-β-Glucanase Preparation against Reference PolysaccharidesSDS–PAGE of the xyloglucan-specific endo-(1
4)-β-glucanase preparation gave only a single band with an apparent molecular weight of 34 kDa (data not shown), as previously reported by Pauly et al. (1999). Incubation of the preparation with tamarind xyloglucan released oligosaccharides that were detected by MALDI–TOF MS, giving ions with m/z of 1085, 1247, and 1409, corresponding to the [M+Na]+ adduct ions of XXXG (12%), XLXG/XXLG (52%), and XLLG (36%), respectively. No oligosaccharides were released from amylose, amylopectin, arabinoxylan, or glucomannan. However, oligosaccharides were released from the (1 3), (1 4)-β-glucan, indicating the presence of some endo-β-glucanase activity that is not xyloglucan specific. Analysis by MALDI–TOF MS gave ions with m/z of 527 (42%), 689 (24%), 851 (12%), 1013 (10%), 1175 (6%), 1338 (2%), 1500 (1%), 1662 (1%), and 1824 (1%), corresponding to the [M+Na]+ adduct ions of oligosaccharides containing 3–11 glucosyl residues. Nevertheless, the proportions of xyloglucan oligosaccharides released by the xyloglucan-specific endo-(1 4)-β-glucanase preparation and a pure cellulase preparation from the 6-M NaOH extract of Vigna radiata were similar as determined by HPAEC (Table 1), indicating that the oligosaccharides released by the xyloglucan-specific endo-(1 4)-β-glucanase were not ‘trimmed’ by other enzymatic activities in the preparation.
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Xyloglucan Oligosaccharides Released from 6-M NaOH Extracts of Alcohol-Insoluble Residues of Monocotyledon Species
A mixture of xyloglucan oligosaccharides obtained by mixing equal volumes of xyloglucan-specific endo-(1
4)-β-glucanase hydrolysates of the 6-M NaOH extracts of Vigna radiata and Nicotiana tabacum was used as a reference mixture for HPAEC. The mixture gave resolved peaks for the oligosaccharides XXG, XXGG, XXXG, XXFG, XLXG, XLFG, XXLG, and XLLG; XSGG was a shoulder on the leading edge of XXXG (Figure 2A). The retention times of the oligosaccharides relative to XXXG are shown in Table 1. The [M+Na]+ adduct ions of the oligosaccharides are shown in Figure 3A and in Table 2.
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The relative proportions of xyloglucan oligosaccharides for a particular species determined by HPAEC (Table 1) and MALDI–TOF MS (Table 2) were similar. Furthermore, there were few differences in the proportions of xyloglucan oligosaccharides from walls of the same species and organ, but at different stages of development (see results for Flagellaria indica in Tables 1 and 2), although there were slightly greater differences in proportions obtained from walls of the same species, but from different organs (see results for Hedychium greenii in Tables 1 and 2). The profiles of xyloglucan oligosaccharides from the non-commelinid monocotyledons were, with the exception of Lemna minor, similar to those from the eudicotyledons Vigna radiata and Arabidopsis thaliana. XXXG was the most abundant xyloglucan oligosaccharide and the fucosylated oligosaccharides XXFG and XLFG were present, although in variable amounts (Tables 1 and 2). XXXG was the predominant core motif, accounting for >92% of the units estimated by HPAEC (Table 1). The chromatogram and mass spectrum of Alstroemeria aurantiaca (Alstroemeriaceae) hydrolysate are shown in Figures 2B and 3B, respectively. The xyloglucan oligosaccharide profile of L. minor was quite different. XXG was the most abundant oligosaccharide, the fucosylated oligosaccharides XXFG and XLFG were not detected, and XXGn was the predominant core motif (
67% by HPAEC) (Tables 1 and 2). In addition, this species gave a range of large unidentified peaks (Table 1) and ions with m/z of 821, 983, 1145, and 1439 (Table 2) corresponding to [M+Na]+ adduct ions of Hex4Pent1, Hex5Pent1, Hex6Pent1, and Hex7Pent2, respectively. These ions may arise from XGGG, XGGGG, XGGGGG, and XXGGGGG (or isomers of these), respectively. In the commelinid monocotyledons, Phoenix canariensis (Arecaceae) had a similar xyloglucan oligosaccharide profile (Figures 2C and 3C, and Tables 1 and 2) to those of the eudicotyledons V. radiata and A. thaliana and the non-commelinid monocotyledons except for L. minor. Dasypogon obliquifolius, in the unplaced family Dasypogonacecae, also had a similar xyloglucan oligosaccharide profile (Figures 2D and 3D). However, the other commelinid taxa had quite different profiles.
Outside the Poales, all species in the Zingiberales and Commelinales had similar profiles, with both XXGn and XXXG core motifs present, with the proportion of XXGn usually greater than XXXG. XXFG was detected in all species by HPAEC and in most species by MALDI–TOF MS, but XLFG was not detected using either technique (Tables 1 and 2). The chromatogram and mass spectrum of the Eichhornia crassipes (Pontederiaceae) hydrolysate are shown in Figures 2E and 3E, respectively. In addition, chromatograms of hydrolysates from all species examined contained an unknown component with a retention time relative to XXXG of 0.96–0.98, which formed a shoulder on the leading edge of the XXXG peak (Figure 2E and Table 1). The proportions of this unknown component varied among the different species, varying from a trace in Maranta leuconeura to 20% in Zingber officinale (Table 1). It is possible that this shoulder is XSGG. It had the same retention time as the XSGG in the Nicotiana tabacum hydrolysate and the m/z of its sodium adduct ion, 1085, is identical to that of XXXG. Hydrolysates of several of the species in the Zingiberales and Commelinales gave mass spectra with abundant proportions of the ions m/z 659 and smaller proportions of m/z 821 (Table 2), corresponding to [M+Na]+ adduct ions of Hex3Pent1 and Hex4Pent1, respectively. These ions may be from XGG and XGGG (or isomers of these).
Within the Poales, there was considerable variation in xyloglucan oligosaccharide profiles. The two species of Bromeliaceae examined, Ananas comosus and Aechmea fasciata, both had XXXG and XXGn core motifs, with XXXG 69 and 40%, respectively, by HPAEC, and 52 and 19%, respectively, by MALDI–TOF MS. The chromatogram and spectrum for the A. comosus hydrolysate are shown in Figures 2F and 3F, respectively. Both species had XXFG, but A. comosus had a much higher proportion (33 and 21% by HPAEC and MALDI–TOF MS, respectively) than A. fasciata (2% by both methods). However, A. fasciata had no XLFG and only small proportions were found in A. comosus (<1 and 3% by HPAEC and MALDI–TOF MS, respectively). Both of these Bromeliaceae species had ions with m/z of 659 and 821 in their mass spectra (see above).
The xyloglucan oligosaccharide profiles of Flagellaria indica (Flagellariaceae) also contained both XXXG and XXGn core motifs (67–68% XXXG by HPAEC and 45–47% by MALDI–TOF MS), with high proportions of XXXG, XXLG, and XXG; XXFG was present but XLFG was not (Figures 2G and 3G). However, both F. indica samples showed a slight shoulder in the HPAEC at the retention time of 0.97 relative to XXXG (see above) (Table 1). In preliminary experiments in which 6-M NaOH extracts of F. indica and Poaceae species had not been treated with lichenase before treatment with the xyloglucan-specific endo-(1 4)-β-glucanase preparation, the chromatograms and mass spectra showed peaks and ions, respectively, corresponding to glucosyl oligosaccharides released from (1 3), (1 4)-β-glucan. However, these peaks and ions were absent when 6-M NaOH extracts were used that had been lichenase pretreated.
In the Poaceae, the XXGn core motif predominated, with Lolium multiflorum, L. perenne, Festuca arudinacea, and Triticum aestivum all having only this core motif (Tables 1 and 2). However, the other three Poaceae species had small proportions of xyloglucan oligosaccharides that probably had the XXXG core motif; these oligosaccharides were more apparent by HPAEC than by MALDI–TOF MS. The chromatogram and mass spectrum of the F. arundinacea hydrolysate are shown in Figures 2H and 3H, respectively. All Poaceae species had a peak with a retention time of 0.95–0.97 relative to XXXG, which was possibly XSGG (see above). XLFG and XXFG were not detected by either technique in any Poaceae species. In addition, the ion with m/z 659 corresponding to the [M+Na]+ adduct ion of Hex3Pent1, possibly XGG or an isomer of this (see above), was present in the spectra of hydrolysates of all Poaceae species except Avena sativa and L. multiflorum (Table 2).
In the other Poales species examined, in the families Restionaceae, Xyridaceae, Cyperaceae, Juncaceae, and Typhaceae, the xyloglucan oligosaccharide profiles showed that the core motif was predominantly XXXG (99% by HPAEC and 100% by MALDI–TOF MS), with the most abundant oligosacccharide being XXXG; XXFG was present, but not XLFG (Tables 1 and 2). The chromatogram and mass spectrum of the Elegia capensis (Restionaceae) hydrolysate are shown in Figures 2I and 3I, respectively.
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DISCUSSION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Monocotyledons Have Xyloglucans with Diverse Structures
Our survey showed that Poaceae are not unique among the monocotyledons in having xyloglucans that are structurally different from the fucogalactoxyloglucans found in the primary walls of most eudicotyledons. Variation in xyloglucan structure is particularly apparent in the commelinid monocotyledons. This structural variation is summarized in Figure 4 in the context of monocotyledon phylogeny based on nucleotide sequences of the plastid genes atpB, matK, ndhF, and rbcL, the mitochondrial gene atp1, and the nuclear genes 18S and partial 26S rDNA (Chase et al., 2006). Support was low for the position on the tree of the family Dasypogonaceae, which was not placed in an order in the classification of APG II (2003), and its xyloglucan structure will be discussed separately (see below).
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Poaceae Xyloglucans
Our results are consistent with other structural studies of Poaceae xyloglucans in showing that XXGn was the predominant repeating core motif and no fucosylated units were present. In an earlier study of the xyloglucans in the walls of barley (Hordeum vulgare) coleoptiles, Gibeaut et al. (2005) also used a xyloglucan-specific endo-(1
4)-β-glucanase and analyzed the xyloglucan oligosaccharides released using MALDI–TOF MS, but treated the wall preparation directly with the enzyme, resulting in acetylated, as well as non-acetylated, oligosaccharides being obtained. The most abundant oligosaccharide was XXGGG, but large proportions of XXGG and XXGGGG were also found. In an early study, Kato et al. (1981) also concluded that the predominant repeating unit of barley seedling xyloglucan was XXGGG. The larger proportions of XXG and smaller proportions of XXGGG and XXGGGG found in the present study probably resulted from the removal, by the alkali extraction, of acetyl groups, which probably occur on glucosyl residues without attached xylosyl residues, except for the reducing glucosyl residue (Figure 1) (Gibeaut et al., 2005). The resulting series of unsubstituted glucosyl residues would be susceptible to glucanase hydrolysis. Although Gibeaut et al. (2005) did not report the presence of galactosyl residues in barley xyloglucan, XLGG found in the present study would give the same m/z sodium adduct ion as XXGGG and so may have been present. Kato et al. (2004a) also reported the presence of galactosyl residues in barley seedling xyloglucans, including LXGG, XLGGG, LXGGG, XLGGGG, and LXGGGG. Our results also indicated that barley, maize (Zea mays), and oat (Avena sativa) walls released small proportions of oligosaccharides with the XXXG core motif; some of these oligosaccharides contained galactosyl residues. The release of XXXG and a galactosylated XXXG by cellulase has previously been reported from a xyloglucan from the walls of suspension cultured rice (Oryza sativa) cells (Kato and Matsuda, 1985). The release of small amounts of XXXGG was also reported by Gibeaut et al. (2005) in their study of barley coleoptile walls.
Small proportions of fucosyl residues, together with the XXXG core motif, have also been detected in a Poaceae xyloglucan using radioactive labeling (McDougall and Fry, 1994). Suspension-cultured cells of tall fescue (Festuca arundinacea) were incubated with exogenous L-[1-3H]fucose, which is known to be incorporated into only fucose and methyl fucose. Extracted xyloglucans were treated with cellulase and radioactive XXFG was detected. As the authors indicated, this finding showed that this species at least has the genetic capacity to synthesize fucosylated xyloglucans. However, we found no evidence of fucosylated xyloglucan oligosaccharides in this, or any other, species of Poaceae examined.
Other Monocotyledon Xyloglucans and their Evolution
As in the primary walls of most eudicotyledons, the xyloglucans of the non-commelinid monocotyledons we examined, with only one exception, Lemna minor (Araceae), were fucogalactoxyloglucans with a XXXG core motif and containing high proportions of XXFG and XLFG. Our results for the structure of onion (Allium cepa) xyloglucan were similar to those reported previously (Ohsumi and Hayashi, 1994). It is likely that during the evolution of non-commelinid monocotyledons from more basal angiosperm taxa, the fucogalactoxyloglucan structure was conserved.
In the same way, this xyloglucan structure was conserved during the evolution of the Arecales (palms) that form the most basal order of the commelinid monocotyledons (Figure 4). However, during the evolution of commelinid monocotyledons more derived than the Arecales, considerable changes occurred in the xyloglucan structures. The major sister clades Zingiberales and Commelinales (Figure 4) have xyloglucans with similar structures, which differ from fucogalactoxyloglucans in several respects: they have XXGn as well as XXXG core motifs, and they have small proportions of XXFG units, but XLFG units are absent. Thus, during the evolution of these orders, XLFG units in the xyloglucans were lost and the XXGn core motif was gained.
Similar changes occurred during the evolution of the basal Poales family Bromeliaceae. In the xyloglucans of this family, both types of core motifs are present; XXFG is present, but XLFG is either absent or present in only very small proportions (Table 1 and Figure 4). Pineapple (Ananas comosus) was found not to have fucogalactoxyloglucans as had previously been reported (Kato et al., 2001). The xyloglucans of the remaining Poales families that have been analyzed, except Flagellariaceae and Poaceae, have similar structures to each other: they have only the XXXG core motif and XXFG units, but no XLFG units are present (Figure 4). There is thus no change in the two fucosylated oligosaccharide units, but, oddly, the phylogeny indicates a reversion to only the XXXG core motif.
The final changes in xyloglucan structure occured in the Flagellariaceae and Poaceae, with the xyloglucans of Flagellaria indica being intermediate in structure between those of the Poaceae and the Poales families Typhaceae to Restionaceae (Figure 4). During the evolution of F. indica xyloglucans, the XXGn core motif was apparently regained, and in the transition to the Poaceae, almost all units with the XXXG core motif were lost as well as all fucosylated units (see above).
As indicated above, the phylogenetic position of the family Dasypogonaceae is uncertain and Chase et al. (2006) commented that it could be finally placed in either the Poales or Arecales (palms). In the Poales, its position varies. In Figure 4, it is sister to the rest of the Poales (58% bootstrap support), but in a study using only 26S rDNA sequences, it is sister to Anarthriaceae (68% bootstrap support) (Neyland, 2002). The xyloglucan oligosaccharide profile of Dasypogon obliquifolius is consistent with it being a fucogalactoxyloglucan, as found in the walls of the Arecales (palms). However, the proportion of XLFG units is lower than in many fucogalactoxyloglucans and the xyloglucans of more species in the Dasypogonaceae need to be examined.
Xyloglucan Variation and Non-Cellulosic Polysaccharide Compositions
Evolution of variation in monocotyledon xyloglucan structure is accompanied by variation in the proportions and structures of the other primary wall non-cellulosic polysaccharides. Both types of variation occur to the greatest extent in the commelinids. The fucogalactoxyloglucans that occur in the primary walls of most non-commelinid monocotyledons and the commelinid order Arecales (palms) are accompanied by large proportions of pectic polysaccharides (Harris et al., 1997; Harris, 2005). However, the xyloglucans in the non-commelinid aquatic plant Lemna minor (duckweed) (Araceae although formerly in the family Lemnaceae) gave quite a different oligosaccharide profile from that of the fucogalactoxyloglucans, including significant proportions of oligosaccharides with the XXGn core motif, neither of the fucosylated oligosaccharides XXFG and XLFG, and many unidentified oligosaccharides. This contrasted with the fucogalactoxyloglucan-like oligosaccharide profiles obtained from the walls of Zantedeschia aethiopica, also in the Araceae. The walls of L. minor, as well as those of a number of other aquatic species in the Alismatales, contain large proportions of pectic polysaccharides that are unusual in containing the apiogalacturonan domain composed of a galacturonan backbone bearing mono- and di-apiosyl side chains (Harris, 2005). The presence of this domain may be related to the aquatic habitat of these species. It would be interesting to know if xyloglucans similar to those in L. minor also occur in the walls of other aquatic species in the Alismatales. If so, these features may be related to the aquatic habitat.
Unlike the primary walls of the non-commelinid monocotyledons, those of the commelinid monocotyledons contain ferulic acid ester-linked to glucuronoarabinoxylans (Smith and Harris, 1995, 2001; Harris et al., 1997; Harris, 2005). However, only small proportions of feruloylated GAXs are present in the primary walls of the basal order Arecales, which have overall non-cellulosic polysaccharide compositions similar to those of the non-commelinid monocotyledons as well as fucogalactoxyloglucans (Carnachan and Harris, 2000).
In contrast, the primary walls of the Zingiberales, Commelinales, and Poales all contain, in addition to pectic polysaccharides and xyloglucans, substantial proportions of feruloylated glucuronoarabinoxylans (Smith and Harris, 1995, 2001; Harris et al., 1997). The primary walls of species of families in the graminoid clade of the Poales (Figure 4) that have been examined have a particularly low content of pectic polysaccharides (Harris et al., 1997; Smith and Harris, 1999). The walls of other families of the Poales and of the Zingiberales and Commelinales have intermediate proportions of pectic polysaccharides (Harris et al., 1997). The primary walls of all the families examined of the Poales, except for the basal families of Bromeliaceae, Typhaceae, and Sparganiaceae, also contain small proportions of (1 3), (1 4)-β-glucans (Smith and Harris, 1999; Trethewey et al., 2005). The proportions of these polysaccharides are highest in the walls of the related families Anarthriaceae, Centrolepidaceae, Ecdeiocoleaceae, Flagellariaceae, and Poaceae.
How the particular variations in structures of monocotyledon xyloglucans found in the present study may be related to their functions is largely unknown, as are the possible functional relationships between variations in xyloglucan structures and variations in the proportions and structures of the other primary wall non-cellulosic polysaccharides. In the Solanaceae and related families that do not have fucogalactoxyloglucans in their primary walls, there is apparently no evidence for variations in the proportions and structures of non-cellulosic polysaccharides. Nevertheless, there have been studies aimed at understanding the possible relationships between xyloglucan structures and their functions. Most of these concern the interactions of xyloglucans with cellulose. For example, xyloglucans with decreased proportions of xylose residues were found to have increased abilities to bind to cellulose in vitro (Chambat et al., 2005). This suggests that xyloglucans with the XXGn core motif bind more readily to cellulose than do xyloglucans with the XXXG repeating core motif. With the latter xyloglucans, similar in vitro experiments indicated that fucosylation assisted with binding to cellulose (Levy et al., 1997). However, studies of Arabidopsis thaliana plants with mutations that affect xyloglucan structure showed that the absence of fucosyl residues on xyloglucans did not affect mechanical strength, although the absence of galactosyl residues did cause some decrease (Ryden et al., 2003; Peña et al., 2004). In addition to interacting with cellulose, there is evidence that substantial proportions of the xyloglucans in primary walls are covalently bound to the pectic polysaccharide rhamnogalacturonan-1 (Thompson and Fry, 2000; Abdel-Massih et al., 2003; Popper and Fry, 2005, 2008). Such interactions between xyloglucans and pectin have been demonstrated in the walls of the Poaceae (maize and barley) and Solanaceae (tomato, Solanum lycopersicum), as well as a range of other eudicotyledon species, indicating that xyloglucans other than fucogalactoxyloglucans are involved. Covalent bonds may also occur between xyloglucans and (1 3), (1 4)-β-glucans in primary walls that contain the latter polysaccharides. Although such bonds have not been found in muro, a highly purified xyloglucan xyloglucosyl transferase from barley (HvXET5) has been shown to catalyze the in vitro formation of covalent linkages between xyloglucans and (1 3), (1 4)-β-glucans (Hrmova et al., 2007). In these experiments, tamarind xyloglucan was used, but it would be interesting to know whether the enzyme is more effective with Poaceae xyloglucans.
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Plant Material
The sources of plant material are shown in Table 3. The orders, families, and species are arranged in alphabetical order according to the classification of APG II (2003). Seedlings of Hordeum vulgare, Zea mays, Triticum aestivum, and Avena sativa were obtained by surface sterilizing the grains in 1% (w/v) NaOCl for 5 min, washing with water, and germinating them on damp vermiculite at 27°C in the dark for 3–7 d. The seedlings were exposed to red light for 1 h daily to prevent mesocotyl elongation. Seedlings of Vigna radiata were obtained in a similar way, except the seeds were surface sterilized in 3% (w/v) NaOCl for 7 h and germinated at 25°C for 4 d. Seedlings of Arabidopsis thaliana were obtained by surface sterilizing in 3% (w/v) NaOCl for 30 min, washing in water, keeping on wet filter paper at 4°C in the dark for 3 d, and germinating in potting mix in an unheated glasshouse (minimum and maximum temperatures 13°C and 27°C, respectively) for 3 weeks. Seedlings of Festuca arundinacea, Lolium multiflorum, L. perenne, and Nicotiana tabacum were obtained by germinating them directly in potting mix in the same glasshouse for 2 or 6 (N. tabacum) weeks.
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Isolation of Alcohol-Insoluble Residues Using 6 M NaOH
Alcohol-insoluble residues were isolated either from intact plant organs that contained mostly non-lignified cell walls or from organs from which tissues containing cells with lignified walls had been removed by peeling or cutting away (Table 3). Lignified cell walls were detected histochemically using bright-field microscopy of fresh transverse sections (cut by hand using a razor blade) by the red color reaction given by phloroglucinol–HCl (Harris et al., 1980). Alcohol-insoluble residues were isolated by a variation of the procedure described by Popper et al. (2001). Plant material (20 g fresh weight) was homogenized in 70% aqueous ethanol (50 mL at 4°C) using a Polytron blender (Model PT10-35, Kinematica GmbH, Luzern, Switzerland) on full power (six 20-s bursts). The homogenate was filtered onto Miracloth (Calbiochem, Darmstadt, Germany) and the residue transferred to a ceramic pestle and mortar containing liquid nitrogen and ground to a fine powder. This powder was incubated in 70% ethanol (150 mL at 60°C) for 6 h to remove soluble sugars. The suspension was filtered onto fresh Miracloth and the residue washed successively with 70% ethanol (100 mL) and acetone (100 mL), dried in a stream of air and stored under vacuum over silica gel.
Extraction of Non-Cellulosic Polysaccharides from Alcohol-Insoluble Residues Using 6 M NaOH
Alcohol-insoluble residues (70–100 mg) were extracted, on an orbital shaker, for 16 h at 37°C with 6 M NaOH containing 1% NaBH4 (10 mL) to prevent alkaline-peeling. The suspension was filtered onto Miracloth, the filtrate cooled on ice for 20 min before its pH was adjusted to 5.0 by adding 17.4 M acetic acid, dropwise. To stop microbial growth, a drop of toluene was added to the neutralized solution, which was then dialyzed at room temperature for 24 h against tap water and then for 1 h against distilled water; the dialysate was freeze-dried.
SDS–PAGE and Specificity of the Xyloglucan-Specific Endo-(1 4)-β-Glucanase Preparation
A freeze-dried preparation of xyloglucan-specific endo-(1 4)-β-glucanase from Aspergillus aculeatus (EC 3.2.1.151
[EC]
) (336 U mg–1), provided by Dr Kirk M. Schnorr, Novozymes, Bagsvaerd, Denmark, was obtained by heterologously expressing a cDNA clone encoding the enzyme in Aspergillus oryzae (Pauly et al., 1999) and was not further purified. The xyloglucan-specific endo-(1 4)-β-glucanase preparation was subjected to SDS–PAGE by the method of Laemmli (1970) using a mini-vertical unit (Hoefer SE 260, Amersham Pharmacia Biotech Europe, GmbH, Freiburg, Germany) and stained with Coomassie Blue.
The specificity of the preparation was determined by incubating reference polysaccharides (2 mg) with the preparation as described below and examining the hydrolysates for oligosaccharides by MALDI–TOF MS. The reference polysaccharides were amylose from potato (Sigma, St Louis, MO, USA), amylopectin from potato (Sigma), xyloglucan from tamarind (Megazyme), (1 3), (1 4)-β-glucan from barley (Megazyme), arabinoxylan from wheat flour (medium viscosity) (Megazyme), and glucomannan from konjac (Amorphophallus konjac) (native, low viscosity) (Megazyme). As controls, each polysaccharide was also incubated with buffer only.
Treatment of Extracted Non-Cellulosic Polysaccharides with Xyloglucan-Specific Endo-(1 4)-β-Glucanase
Polysaccharides extracted with 6 M NaOH from alcohol-insoluble residues (2 mg) were incubated with 0.1% (w/v) xyloglucan-specific endo-(1 4)-β-glucanase in 50 mM ammonium formate buffer (pH 3.0) (0.5 mL) for 5 min at 37°C and stopped by heating to 100°C for 2 min. Because significant proportions of (1 3), (1 4)-β-glucans are known to occur in the primary cell walls of Poaceae and Flagellariaceae (Smith and Harris, 1999; Trethewey et al., 2005) and preliminary experiments had shown that the xyloglucan-specific endo-(1 4)-β-glucanase preparation was able to hydrolyze (1 3), (1 4)-β-glucans, these polysaccharides were removed from the 6-M NaOH extracts of all Poaceae species and Flagellaria indica before they were treated with xyloglucan-specific endo-(1 4)-β-glucanase. This was done by treating the dry extracts (10 mg) with (1 3), (1 4)-β-glucanase (lichenase) from Bacillus subtilis (Megazyme International Ireland Ltd, County Wicklow, Ireland) (EC 3.2.1.73
[EC]
) (40 µL, 2 enzyme units, 1 h at 50°C) in 20 mM sodium phosphate buffer (pH 6.5) (0.8 mL). Water (4.12 mL) was added with a drop of toluene and the solution dialyzed against tap water for 24 h, then against distilled water for 1 h, and freeze-dried. Dry 6-M NaOH extracts (2 mg) of V. radiata were also incubated with an electrophoretically homogeneous endo-(1 4)-β-glucanase (cellulase) preparation (5 enzyme units) from Trichoderma longibrachiatum (Megazyme) in 50 mM ammonium formate buffer (pH 5.0) (0.2 mL) for 16 h at 37°C and the reaction stopped as described above. (This cellulase preparation is stated by Megazyme to also have activity towards birchwood xylan and, in the present study, we found it had activity towards arabinoxylan from wheat flour.) Control, buffer-only incubations were also carried out using the conditions for the xyloglucan-specific endo-(1 4)-β-glucanase and the cellulase.
High-Performance Anion-Exchange Chromatography (HPAEC)
Xyloglucan oligosaccharides were separated and detected using a Dionex BioLC system (Dionex, Sunnyvale, CA, USA) fitted with a pulsed amperometric detector. They were separated at 28°C on a CarboPac PA1 analytical anion-exchange column (250 x 4 mm) fitted with a CarboPac PA1 guard column (50 x 4 mm) using a linear gradient from 50 mM NaOAc in 100 mM NaOH (solvent A) to 100 mM NaOAc in 100 mM NaOH (solvent B) over 110 min at 1 mL min–1. After each run, the column was washed for 10 min with solvent B, 5 min with 300 mM NaOH, and 30 min with solvent A. Peak areas on chromatograms were quantified by cutting out and weighing. After running each unknown sample, a reference standard of xyloglucan oligosaccharides from Vigna radiata and Nicotiana tabacum (see below) was run.
Chromatogram peak areas were quantified by cutting out and weighing. The oligosaccharides XXXG and XXGGG co-chromatographed, but gave ions of m/z 1085 and 1115, respectively, by MALDI–TOF MS. The relative contribution of the two oligosaccharides to the chromatogram peak was estimated from the relative strengths of these ions. However, because XXGGG has the same molecular weight as XLGG, the strength of the 1115-m/z ion was first corrected for the XLGG contribution using the peak area due to this oligosaccharide on the chromatogram.
Matrix-Assisted Laser-Desorption Ionization Time-of-Flight Mass Spectrometry (MALDI–TOF MS)
The molecular weights of the sodium adducts of oligosaccharides [M+Na]+ were determined using a Voyager-DETM PRO MALDI–TOF Workstation (Applied Biosystems, Foster City, CA, USA). The xyloglucan-specific endo-(1 4)-β-glucanase hydrolysate (10 µL) or reference tamarind XG oligosaccharide preparations (Megazyme) (10 µL) (XXXG 1.06 mg mL–1 in water; a mixture of XXXG, XLXG, XXLG, and XLLG 3.67 mg mL–1 in water) was mixed with 2,5-dihydroxybenzoic acid (Sigma) (10 µL, 10 mg mL–1 in water) and NaCl (6 µL, 10 mM in water). The mixture (1 µL) was dried onto a sample plate, and the spectrometer operated in the reflectron mode at an accelerating voltage of 20 kV with a delay time of 200 ns. Each spectrum consisted of data from 100 laser shots.
Reference Oligosaccharides and Assignment of Peaks
A range of mixtures of reference xyloglucan oligosaccharides was used to determine relative retention times by HPAEC. All of these mixtures were also subjected to MALDI–TOF MS.
The oligosaccharide XXXG and a mixture of the oligosaccharides XXXG, XLXG, XXLG, and XLLG, obtained by treating tamarind xyloglucan with an endo-glucanase, were from Megazyme. The order of elution of the XLXG, XXLG, and XLLG (Table 1) was assigned by comparing the results with those of Ren et al. (2005), who used similar chromatographic conditions to separate these oligosaccharides. A xyloglucan-specific endo-(1 4)-β-glucanase hydrolysate of the 6-M NaOH extracted polysaccharides from Vigna radiata was used as a reference oligosaccharide mixture, as the xyloglucans of this species are known to yield XXFG and XLFG in addition to the oligosaccharides from tamarind xyloglucan (Kato et al., 2004b). The order of elution of XXFG and XLFG (Table 1) was assigned by comparing the results with those of Lerouxel et al. (2002), who separated Arabidopsis thaliana xyloglucan oligosaccharides, which are the same as those of Vigna radiata (Table 1).
A xyloglucan-specific endo-(1 4)-β-glucanase hydrolysate of the 6-M NaOH extract from Nicotiana tabacum was also used as a reference oligosaccharide mixture, as the xyloglucans of this species are known to yield XXG, XXGG, and XSGG (Mori et al., 1980; Hoffman et al., 2005). The order of elution of these oligosaccharides (Table 1) was assigned by separating them by gel permeation chromatography and running them individually by HPAEC. Gel permeation chromatography was done using a BioGel P-2 column (fitted with a water jacket held at 50°C) at a flow rate of 0.2 mL min–1 and 120 (1-mL) fractions collected. The phenol-sulfuric acid assay for carbohydrates (Dubois et al., 1956) was used to determine which fractions contained oligosaccharides and these were analyzed by HPAEC and MALDI–TOF MS. Fractions 41, 48, and 54 contained oligosaccharides tentatively identified as XSGG, XXGG, and XXG, respectively, on the basis of MALDI–TOF MS (data not shown). Although the MALDI–TOF MS results were consistent with these oligosaccharides, their sequences are unknown and isomers may have been present, for example, GXXG rather than XXGG (Hoffman et al., 2005). Gel permeation chromatography was also used to assign the retention time (Table 1) of what was tentatively identified by MALDI–TOF MS as XXGGG from a xyloglucan-specific endo-(1 4)-β-glucanase hydrolysate of the 6-M NaOH extract from Hordeum vulgare. Fraction 31 contained mostly XXGGG, but also some XXXG (which co-eluted by HPAEC) (data not shown). Again, the oligosaccharide was not sequenced and so it may be an isomer, such as GXXGG or LXGG (Hoffman et al., 2005).
A reference standard mixture containing XXG, XXGG, XXXG, XXFG, XLXG, XLFG, XXLG, XLLG, and XSGG was obtained by mixing equal volumes of xyloglucan-specific endo-(1 4)-β-glucanase hydrolysates of 6-M NaOH extract Vigna radiata and Nicotiana tabacum, and was run by HPAEC after each unknown sample.
An oligosaccharide with a retention time of 0.76 relative to XXXG was tentatively identified as XLGG by treating an xyloglucan-specific endo-(1 4)-β-glucanase hydrolysate of a 6-M NaOH extract of a primary wall preparation from Musa sp. with β-galactosidase. The peak disappeared and there was a corresponding increase in the size of the XXGG peak (Trethewey et al., in preparation). An ion with m/z of 1277 was also tentatively identified as the [M+Na]+ adduct ion of XXGGGG, but no HPAEC peak was assigned to this oligosaccharide.
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FUNDING |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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The work was supported by funding from the University of Auckland.
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Acknowledgements |
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We thank J. Hsieh for optimizing the MALDI–TOF MS conditions during the tenure of a summer studentship at the School of Biological Sciences, University of Auckland, Professor S.C. Fry and Dr Z.A. Popper, University of Edinburgh, and E.K. Cameron, Auckland War Memorial Museum, for helpful advice and discussion, Dr K.M. Schnorr, Novozymes, Denmark, for the gift of the xyloglucan-specific endo-(1
4)-β-glucanase, and M. Paxton for critically reading the manuscript. No conflict of interest declared. |
Notes |
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a See Figure 1 for the structural nomenclature of xyloglucans.
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