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Molecular genetics of fructan metabolism in perennial ryegrass


  • Jaye Chalmers,

    1. Plant Biotechnology Centre, Primary Industries Research Victoria, Department of Primary Industries and Molecular Plant Breeding CRC, La Trobe University, Victoria 3086, Australia
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  • Angela Lidgett,

    1. Plant Biotechnology Centre, Primary Industries Research Victoria, Department of Primary Industries and Molecular Plant Breeding CRC, La Trobe University, Victoria 3086, Australia
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  • Nicholas Cummings,

    1. Plant Biotechnology Centre, Primary Industries Research Victoria, Department of Primary Industries and Molecular Plant Breeding CRC, La Trobe University, Victoria 3086, Australia
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  • Yingying Cao,

    1. Plant Biotechnology Centre, Primary Industries Research Victoria, Department of Primary Industries and Molecular Plant Breeding CRC, La Trobe University, Victoria 3086, Australia
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  • John Forster,

    1. Plant Biotechnology Centre, Primary Industries Research Victoria, Department of Primary Industries and Molecular Plant Breeding CRC, La Trobe University, Victoria 3086, Australia
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  • German Spangenberg

    Corresponding author
    1. Plant Biotechnology Centre, Primary Industries Research Victoria, Department of Primary Industries and Molecular Plant Breeding CRC, La Trobe University, Victoria 3086, Australia
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* Correspondence (fax +61 39479 3618; e-mail german.spangenberg@dpi.vic.gov.au)


Fructans are the main storage carbohydrates of temperate grasses, sustaining regrowth immediately after defoliation, as well as contributing to the nutritive value of feed. Fructan metabolism is based on the substrate sucrose and involves fructosyltransferases (FTs) for biosynthesis and fructan exohydrolases (FEHs) for degradation. Sucrose is also utilized by invertases (INVs), which hydrolyse it into its constituent monosaccharides for use in metabolism. The isolation, molecular characterization, functional analysis, and phylogenetic relationships of genes encoding FTs, FEHs, and INVs from temperate grasses are reviewed, with an emphasis on perennial ryegrass (Lolium perenne L.). The roles these enzymes play in fructan accumulation and remobilization, and future biotechnological applications in molecular plant breeding are discussed.


Fructans are present in 15% of angiosperm species, accounting for nearly one-third of global vegetation cover, and are particularly widespread in the grasses (Chatterton et al., 1989; Hendry, 1993). Fructans are products of the polymerization of sucrose to generate branched or linear polyfructose molecules that accumulate in plants in addition to or instead of starch (Hendry, 1993). Plant fructans are highly accessible storage carbohydrates that are sequestered in the leaf base and storage organs and are mobilized when photosynthetic carbon supply is lower than the demand for growth. Fructans have been implicated in facilitation of sucrose unloading from the phloem and maintenance of osmotic potential to ensure cell enlargement in the elongation zone (Pavis et al., 2001b). Accumulation of fructans in plants has also been associated with tolerance to cold and drought (Hendry, 1993; Pilon-Smits et al., 1995; Konstantinova et al., 2002; Hisano et al., 2004a), particularly in the development of freezing tolerance in cereals (Olien, 1984; Pontis, 1989; Tognetti et al., 1990; Yoshida et al., 1997; Livingston and Henson, 1998; Yoshida et al., 1998; Kawakami and Yoshida, 2002).

Fructans accumulate during periods when supply of assimilated carbon exceeds demand, and decline in association with a transient accumulation of sucrose prior to the onset of rapid spring growth (Thomas and James, 1999). A spatial gradient of carbohydrate content from fructan to hexoses and sucrose is observed from the rapidly elongating tissue at the base of grass leaves and along the longitudinal axis of the elongating blade (Prud’homme et al., 1992). Fructans are predominantly stored in the vacuole in which they are synthesized from sucrose, lowering the sucrose concentration in the cell and preventing sugar-induced feedback inhibition of photosynthesis (Pollock, 1986). An accumulation of sucrose triggers the onset of fructan synthesis (Cairns et al., 1997). In plants that store fructans as their main form of accessible energy reserve, dependency of fructan synthesis on sucrose availability is so high, that fructans have been considered as an extension of the sucrose storage pool (Prud’homme et al., 1992).

The accumulation and remobilization of fructans in plants involves the concerted action of a number of specific fructosyltransferases (FT), fructan exohydrolases (FEH) and invertases (INV) that are tightly and coordinately regulated. Genes encoding these activities have been isolated and characterized (Vijn and Smeekens, 1999; Van Laere and Van den Ende, 2002; Ritsema and Smeekens, 2003a,b). The molecular genetics of fructan metabolism in temperate grasses is reviewed here, with an emphasis on perennial ryegrass (Lolium perenne L.), the predominant grass for global temperate pastoral agriculture.

Fructan biosynthesis

The substrate of fructan biosynthesis is sucrose. Addition of a fructose residue to one of the three primary alcohol groups of sucrose will result in one of the following trisaccharides that are the precursors to all fructans of higher degree of polymerization (DP): 1-kestose, 6-kestose or 6G-kestose (Figure 1) (French, 1989). Fructose–fructose linkages occur between C2 of one fructose residue and the primary alcohol group at C1 or C6 of another forming β(2-1) (inulin) or β(2-6) (levan) linkages, respectively (Pontis and del Campillo, 1985; Suzuki and Pollock, 1986).

Figure 1.

Molecular structures of the three trisaccharide precursors to plant fructans. All structures result from the addition of a fructose residue to sucrose, which consists of a glucose and a fructose residue, and is shown in bold. The β(2-1) and β(2-6) linkages are indicated.

The fructan profiles of plants show significant structural diversity, and are the result of the activity of multiple enzymes. Plant fructans generally show a DP ≤ 50, although some have been shown to exceed DP 200 (Vijn and Smeekens, 1999). Five distinct classes of fructans have been identified in plants: inulin series, levan series, mixed levan (graminan), inulin neoseries, and levan neoseries (Table 1). While dicotyledonous plants predominantly accumulate inulin series fructans (Van Laere and Van den Ende, 2002), the monocotyledonous temperate grasses often produce a mixture of fructan types. L. perenne, for instance, accumulates fructans of the inulin series, inulin neoseries, and levan neoseries (Pavis et al., 2001b).

Table 1.  Summary of the five classes of fructans identified in plants
ClassesLinkage typeSeries trisaccharide
Inulin seriesβ(2-1)1-kestose (G1-2F1-2F)
Levan seriesβ(2-6)6-kestose (G1-2F6-2F)
Mixed levan (graminan)β(2-1) and β(2-6)1-kestose and 6-kestose
Inulin neoseriesβ(2-1)6G-kestose (F2-6G1-2F)
Levan neoseriesβ(2-6)6G-kestose

Two enzymes are required to produce inulin series fructans, the simplest type that is present in plants. The first enzyme, sucrose:sucrose 1-fructosyltransferase (1-SST, EC, transfers a fructose unit from one sucrose molecule to another producing 1-kestose, and a second enzyme, fructan:fructan 1-fructosyltransferase (1-FFT, EC, transfers fructose units from molecules with a DP ≥ 3 on to the growing fructose chain resulting in a complement of fructans with varying chain lengths.

The best-characterized enzyme responsible for the β(2-6) linkages in the levan and mixed levan fructans is sucrose:fructan 6-fructosyltransferase (6-SFT, EC This enzyme sources a fructose unit from sucrose and transfers it on to a wide variety of acceptors including sucrose, 1-kestose and 6-kestose producing 6-kestose, bifurcose (1,6-kestotetraose) or higher DP β(2-6) linked fructans, respectively (Duchateau et al., 1995; Wei et al., 2000). The activity of two other enzymes has been hypothesized to produce β(2-6) linkages in the absence of 6-SFT: fructan:fructan 6-fructosyltransferase (6-FFT) and sucrose:sucrose 6-fructosyltransferase (6-SST). 6-FFT activity has been postulated in L. perenne, and 6-SST activity has been described in crude enzyme extracts from Poa secunda (big bluegrass) (Chatterton and Harrison, 1997; Pavis et al., 2001a; Wei et al., 2002). Although activity of these two enzymes has been predicted for many temperate grasses, the isolation and functional characterization of genes encoding these activities have not yet been reported.

Biosynthesis of neoseries fructans is the result of fructan:fructan 6G-fructosyltransferase (6G-FFT) activity, which catalyses the transfer of a fructose unit from a fructan (e.g. 1-kestose) on to C6 of the glucose unit of another fructan or sucrose (Shiomi, 1989), which is further elongated with β(2-1) or β(2-6) linkages to produce the inulin neoseries or levan neoseries fructans, respectively. 6G-FFT activity from onion, and to a much lesser extent in asparagus, has been shown to produce inulin series and inulin neoseries fructans, bringing into question the need for 1-FFT activity in all species (Vijn et al., 1997; Ritsema et al., 2003; Fujishima et al., 2005; Ueno et al., 2005). 6G-FFT activity in crude enzyme extracts from L. perenne shows some 1-FFT activity, producing DP4 inulin fructans, significantly smaller than those found in planta (Pavis et al., 2001b). The production of levan series and/or levan neoseries fructans by 6G-FFT has not been reported in crude enzyme extracts from species which accumulate this fructan series.

Fructan degradation

By strict definition, fructan hydrolases (EC, which include exo- and endohydrolases, cleave β(2-1) and β(2-6) linked fructose residues in inulin and/or levan type fructans and sucrose (Webb, 1992). Fructan exohydrolases (FEHs), which are the most physiologically relevant in plants, are reportedly capable of hydrolysing terminal fructose residues of fructans, but not sucrose. FEHs can be specific hydrolases of either β(2-1) linkages (1-FEH) or β(2-6) linkages (6-FEH), or they may have the ability to hydrolyse both linkage types. The fructan-degrading activity of plant FEHs is necessary for the degradation of fructans to sucrose for mobilization, but in addition, FEHs may also be implicated in fructan biosynthesis. For example, FEHs may be involved in the biosynthesis from bifurcose of the trisaccharide precursor to the levan series fructans, 6-kestose (Bancal et al., 1992). In addition, simultaneous activity of fructan biosynthetic and breakdown enzymes during graminan biosynthesis has been reported in wheat stems (Bancal et al., 1992; Bancal et al., 1993; Van den Ende et al., 2003).

Conversely, enzymes involved in fructan biosynthesis have also been implicated in fructan degradation. One example is afforded by 1-FFT activity in Helianthus tuberosus (Jerusalem artichoke). Fructan degradation in this species is proposed to result from the action of both 1-FEH and 1-FFT activity, as 1-FFT reversibly transfers fructosyl units from higher DP oligosaccharides to those with lower DPs (Edelman and Jefford, 1968). Although 6-FFT activity has not yet been described, it has been postulated in L. perenne, in which it could also be involved in fructan degradation by analogy to 1-FFT (Pavis et al., 2001b).

Sucrose degradation

In addition to the fructose released by the activity of FEHs, sucrose can be hydrolysed into its constituent hexoses, glucose and fructose, for entry into metabolism, osmoregulation, cell enlargement, and grain filling, or in response to wounding and cold (Sturm, 1999; Morvan-Bertrand et al., 2001). This is achieved in plants by either sucrose synthase (EC or INV (EC activity (Tymowska-Lalanne and Kreis, 1998; Sturm, 1999). INVs show variation for isoform structure, biochemical properties, and subcellular localizations (Tymowska-Lalanne and Kreis, 1998; Sturm, 1999). There are three main types of INVs in plant cells: vacuolar, extracellular (cell wall) and cytoplasmic. Vacuolar invertases (VINVs) accumulate as soluble proteins in the lumen of the vacuole. Together with cell wall invertases (CWINVs), which are ionically bound to the cell wall, they hydrolyse a variety of fructose-containing oligosaccharides, have a pH optimum between 4.5 and 5, and are thus collectively known as acid INVs. Cytoplasmic INVs prefer a neutral or slightly alkaline pH and are sucrose specific (Sturm, 1999). As fructans are reported to occur predominantly in the vacuoles of plant cells, the acid INVs are believed to be most physiologically relevant to fructan mobilization in plants ( Wagner et al., 1983; Frehner et al., 1984).

VINVs and FTs show high sequence homology and similar enzyme activity. While FTs transfer fructose from sucrose to a sucrose or fructan acceptor, VINVs transfer a fructose unit from sucrose to an acceptor water molecule. VINV activity in the Gramineae is the result of multiple isoforms of this enzyme, with different expression patterns and tissue distributions (Simpson et al., 1991; Bonnett and Simpson, 1993; Walker et al., 1997; Gallagher et al., 2004).

Fructan metabolic pathways

Fructans that accumulate in plants vary in linkage structure, extent of branching, and size distribution, and these characteristics are species-dependent. Although the number of potential plant fructan oligomers is numerous, it appears that the variety of structures present in any species are not random and that a specific linkage pattern is preferred (Pollock and Cairns, 1991; Housley and Pollock, 1993). In general, inulin-type fructans are found in the dicotyledonous and non-graminaceous monocotyledonous plants, whereas all types are found across the temperate grasses (Wei et al., 2000). In those cases in which only inulin-type fructans are present, clear pathways for fructan biosynthesis and degradation can be defined (Van Laere and Van den Ende, 2002). However, in temperate grasses, because of the complexity of fructan profiles found, only hypothetical models for fructan metabolism can be proposed, as shown for L. perenne in Figure 2 (Figure 2).

Figure 2.

Proposed fructan metabolism pathway in Lolium perenne. The substrate of fructan biosynthesis is sucrose, which is utilized by 1-SST (sucrose:sucrose 1-fructosyltransferase) to produce 1-kestose. This structure is then either elongated with β(2-1) linkages by 1-FFT (fructan:fructan 1-fructosyltransferase), producing inulin series fructans, or used by 6G-FFT (6-glucose fructosyltransferase) to produce 6G-kestose, the trisaccharide precursor to the neoseries fructans. 6G-kestose is then either elongated by 1-FFT to produce the inulin neoseries fructans, or by a 6-FFT (fructan:fructan 6-fructosyltransferase) or 6-SFT (sucrose:fructan 6-fructosyltransferase) with β(2-6) linkages to produce levan neoseries fructans.

The temperate grasses Dactylis glomerata (cocksfoot or orchard grass), P. secunda and Phleum pratense (Timothy grass) contain predominantly linear levan-type fructans based on the single trisaccharide precursor 6-kestose (Suzuki and Pollock, 1986; Chatterton et al., 1993; Chatterton and Harrison, 1997). Although their fructan profiles are relatively simple, it is still not clear whether fructan biosynthesis in these species involves a single enzyme like 6-SFT or if specific 6-SST and 6-FFT enzymes are involved.

The most complex fructan profiles identified in plants belong to other members of the Poaceae family, producing fructans with both β(2-1) and β(2-6) linkages in substantial amounts (Pavis et al., 2001b). The factors determining the types of accumulated fructans are not fully understood. Species of the genera Avena and Lolium, which belong to the closely allied Poaceae tribes Aveneae and Poeae, respectively, contain fructans with some internal glucose molecules, while Triticum aestivum (bread wheat) and Bromus tectorum (cheatgrass), which belong to the closely allied Triticeae and Bromeae tribes (Soreng and Davis, 1998) produce fructans with only terminal glucose residues (Bancal et al., 1992; Sims et al., 1992; Chatterton et al., 1993; Livingston et al., 1993). L. temulentum (darnel) accumulates all three trisaccharide precursors and multiple series of fructan oligomers, thus displaying the most complex fructan profile so far identified in plants (Cairns and Pollock, 1988).

The fructan profile of L. perenne includes inulin series, inulin neoseries and levan neoseries fructans (Pavis et al., 2001b). The most abundant trisaccharides present in L. perenne are 1-kestose and 6G-kestose, with 6-kestose present in significantly smaller amounts (Smouter and Simpson, 1991; Sims et al., 1992; Pavis et al., 2001b). Internal glucose residues are present in 75% of fructans with DP ≥ 8 in this species (Pavis et al., 2001b). Based on this profile, it has been proposed that at least four enzymes are required to produce this complement of fructans in L. perenne: 1-SST, 1-FFT, 6G-FFT, and either 6-FFT or 6-SFT (Pavis et al., 2001a). Sucrose is utilized by 1-SST to produce 1-kestose that is either elongated with β(2-1) linkages by 1-FFT to produce inulin series fructans, or is utilized by 6G-FFT to produce 6G-kestose. 1-FFT or 6-FFT/6-SFT elongate 6G-kestose with β(2-1) or β(2-6) linkages to produce inulin neoseries or levan neoseries fructans, respectively (Figure 2). The notable absence of bifurcose, which is usually indicative of 6-SFT activity, may favour the presence of 6-FFT rather than 6-SFT activity. Genes encoding 6-FFT activity have not as yet been described in any target species. The presence of both inulin-and levan-type fructans in L. perenne suggests the presence of separate FEHs with 1-FEH or 6-FEH activity, and/or enzymes showing both types of activity. Purification of 6-FEH activity has been described in L. perenne, providing evidence for FEHs with specific activity (Marx et al., 1997).

Isolation of fructan metabolism genes from L. perenne

Very few full-length cDNA sequences encoding fructan metabolism genes have been isolated from temperate grass species. Most of the sequences identified are only partial in length and tentatively annotated. The most comprehensive set of cDNAs encoding enzymes involved in fructan metabolism that have been isolated and characterized from temperate grasses is from L. perenne, and includes LpFT1 (accession AF481763), LpVINV (LpFT2, AY082350), Lp1-SST (LpFT3, AY245431), Lp1-FFT (AB186920), Lp6G-FFT (AB125218), LpFT4 (DQ073970), LpFEH (DQ073968), and LpCWINV (DQ073969) (Lidgett et al., 2002; Chalmers et al., 2003; Johnson et al., 2003; Hisano et al., 2004b,c). Two additional sequences, LpSST (AF492836) and Lp6-FT (AF494041), which are almost identical to Lp6G-FFT (98.5%) and Lp1-FFT (99.5%), respectively, are available (Lasseur et al., 2002a,b). However, recent functional data suggest LpSST encodes a 6G-FFT (Lp6G-FFT), while Lp6-FT codes for a 1-FFT (Lp1-FFT) (Toshihiko Yamada, personal communication). Genomic sequences for LpFT1, Lp1-SST and LpFT4 have also been isolated (Lidgett et al., 2002; Chalmers et al., 2003).

Sequence analysis

Fructan metabolism enzymes are glycosidases that catalyse the hydrolysis of glycosidic bonds (Pons et al., 1998). More specifically, they belong to glycoside hydrolase family 32 because of their overall amino acid sequence similarity including the conserved active site residues Asp (D) in the sucrose binding box NDPNG and Glu (E) in the conserved block WECXD (Henrissat, 1991). The functions of these conserved residues have been confirmed through site-directed mutagenesis of yeast, bacterial and plant enzymes (Pons et al., 2004). A third active site residue, Asp (D) in FRDP, has been proposed based on homology and site-directed mutagenesis in related enzymes (Pons et al., 2004). The X-ray diffraction structure of an FEH from Cichorium intybus (Ci1-FEHIIa) has recently been resolved and these three conserved regions are shown to cluster in the putative active site of this enzyme (Verhaest et al., 2005). These three regions are conserved in all FT, INV, and FEH deduced amino acid sequences isolated from plants (Figure 3). The FRDP motif only shows variation in the first amino acid, reflecting the pattern [F/Y]RDP. The WECXD motif falls in the middle of a highly conserved region 12 amino acids in length, the core of which displays the pattern [L/V/W/Y]EC[I/L/M/P/V][D/E]. Much more variation is present in the NDPNG motif, which better reflects the pattern HXX[P/T/V]XXXX[A/I/L/M/V][A/C/G/N/S/Y][D/E]P[C/D/N/S][A/G] proposed as a modified sequence pattern to classify glycoside hydrolase family 32 members (Pons et al., 2000). The NDPNG pattern is conserved in the graminaceous monocotyledon 1-SSTs and all VINVs (Figure 3).

Figure 3.

Deduced amino acid sequence alignment of three well conserved regions of FTs, FEHs and INVs from temperate grasses and cereals. Sequence accession numbers are given in Figure 4.

The LpFT1 and LpFT4 genomic sequences contain four exons, including a 9 bp mini exon encoding the core tripeptide (DPN/S) of the conserved motif NDPNG involved in sucrose binding (Sturm, 1999; Lüscher et al., 2000b; Lidgett et al., 2002; Yoshida et al., 2004). The Lp1-SST gene lacks this mini exon, with the conserved tripeptide instead forming the first three amino acids of a larger second exon (Chalmers et al., 2003). This mini exon is also absent in the 1-SST gene (Fa1-SST) from tall fescue (Festuca arundinacea Schreb.) (Lüscher et al., 2000a). It has been identified in the 1-SST gene (Ta1-SST) from wheat (Yoshida et al., 2004). Alternative splicing of the mini exon in L. perenne may play a role in the regulation of endogenous fructan pools under certain physiological conditions, as has been reported for an acid INV from potato under cold stress (Bournay et al., 1996; Lidgett et al., 2002).

Plant fructan metabolism genes generally share high sequence similarity, making it difficult to infer specific putative cellular functions based on deduced amino acid sequence information only. FTs and VINVs are vacuolar-localized, and therefore include a relatively long signal peptide as the vacuolar-targeting signal, while the CWINVs and FEHs contain much shorter signal peptides (Van den Ende et al., 2002). The FTs and VINVs from L. perenne include signal peptides ranging in length from 59 to 115 amino acids, while those of LpCWINV and LpFEH are 22 and 23 amino acids in length, respectively (Table 2). The isoelectric points (pI) of the mature proteins also support the extracellular occurrence of LpCWINV, which has a relatively high pI (8.49), while the remaining enzymes have pIs in the range of 4.67–6.10, consistent with their vacuolar localization (Table 2). FEHs are unusual in that they have short signal peptides in common with extracellular enzymes, and low pIs characteristic of vacuolar enzymes, which are contradictory characteristics in relation to cellular localization. However, as fructans are predominantly found in vacuoles, and FEH activity in H. vulgare leaves has been localized to the vacuole, it is most likely that FEHs are vacuolar enzymes (Wagner and Wiemken, 1986).

Table 2.  Protein properties derived from deduced amino acid sequences of fructan metabolism genes in Lolium perenne. Sequence accession numbers are given in Figure 4. MW molecular weight, pI isoelectric point, Gly glycosylation
NameProtein sizeMW (KDa)Signal peptide (N°. residues)Mature protein pI valueGly sites N-X-S/T
Lp1-FFT62362.5 595.077
Lp6G-FFT64564 634.687
LpCWINV58362 228.494
LpFEH57062 235.987

Phylogenetic analysis suggests that monocotyledonous FTs are more similar to monocotyledonous INVs than dicotyledonous FTs, while dicotyledonous FTs share higher sequence similarity with dicotyledonous INVs than with monocotyledonous FTs (Van der Meer et al., 1998; Wei and Chatterton, 2001). The high sequence homology, conserved gene structure, and biochemical activity generally found in VINVs and FTs, and the relatively distant relationship between CWINVs and VINVs have led to a hypothesis for the evolutionary origin for these enzymes (Wei and Chatterton, 2001). It can be postulated that duplication of an ancestral INV gene occurred before the divergence of monocotyledons and dicotyledons, and that these duplicates became CWINV and VINV isoforms, the latter then diverging again in both lineages to form FTs. Dicotyledon FEHs are most homologous to CWINVs and are therefore likely derived from an ancestral CWINV gene, acquiring a low isoelectric point and a vacuolar-targeting signal (Van den Ende et al., 2002; Van Laere and Van den Ende, 2002). Phylogenetic analysis using monocotyledon sequences exclusively reveals grouping of FTs and VINVs separately from FEHs and CWINVs, consistent with the proposed model for their evolutionary origin (Figure 4). This analysis also reveals very distinct groupings within these two main divisions on the phylogenetic tree, including a close relationship between monocotyledon FEHs and CWINVs, suggesting a similar evolutionary origin to that described for dicotyledonous plants.

Figure 4.

Unrooted phylogenetic dendrogram constructed from monocot fructosyltransferase, acidic vacuolar invertase, and fructan exohydrolase deduced amino acid sequences. Their respective nucleotide sequence accession numbers are: Lolium perenne LpFT4 DQ073970, Hordeum vulgare Hv1-SST AJ567377, T. aestivum Ta1-SST AB029888, L. perenne Lp6G-FFT AB125218, Festuca arundinacea Fa1-SST AJ297369, L. perenne Lp1-SST AY245431, Agropyron cristatum Ac6-SFT AF211253, H. vulgare Hv6-SFT X83233, Triticum aestivum Ta6-SFT AB029887, Poa secunda Ps6-SFT AF192394, L. perenne Lp1-FFT AB186920, L. temulentum Lt6-FT AJ532550, L. perenne LpFT1 AF481763, L. perenne LpVINV AY082350, L. temulentum LtINV1:4 AJ532549, Oryza sativa OsVINV3 AF276704, Saccharum officinarum SoVINV AF062735, O. sativa OsVINV2 AF276703, Zea mays ZmIvr1 U16123, T. aestivum TaCWINV AF030420, L. perenne LpCWINV DQ073969, Z. mays ZmCWINV AF165179, O. sativa OsCWINV AF155121, L. perenne LpFEH DQ073968, H. vulgare Hv1-FEH AJ605333, T. aestivum Ta1-FEH AJ508387. The scale bar indicates a distance value of 0.1.

Comparison of the deduced amino acid sequences of the L. perenne sequences to other available sequences, in combination with phylogenetic analysis has permitted the assignment of putative functions to most of these sequences with some degree of confidence (Table 3, Figures 3 and 4). The deduced amino acid sequences of Lp1-SST, LpVINV, LpFEH, and LpCWINV all show highest sequence identity to, and closest phylogenetic clustering with, other sequences assigned to these respective functions. Lp1-FFT and Lp6G-FFT clearly group with, and show the highest levels of sequence identity to other FTs. The functions of LpFT1 and LpFT4 remain largely unclear, both showing highest levels of sequence identity to FTs, but grouping separately to FTs and VINVs in the phylogenetic analysis.

Table 3.  Deduced amino acid sequence identity (%) of fructan metabolism genes from selected monocotyledons. Sequence accession numbers are given in Figure 4
Fa1-SST 66846665667259636336393843
Lp1-FFT  659767746965626337363840
Lp6G-FFT   6563667063616338383939
Lt6FT    68746965616336353739
Ac6-SFT     686764596237363939
Ps6-SFT      7067626438393841
LpFT1       67687138404045
LpFT4        616037393942
LpVINV         9037383943
LtINV1:4          39394243
LpFEH           743851
Ta1-FEH            3748
LpCWINV             70

Genomic organization of fructan metabolism genes

Determination of gene copy number has not been widely reported for fructan metabolism genes of temperate grasses. Hv6-SFT has been reported to be a single copy gene in Hordeum vulgare L. (barley), while multiple copies of Ac6-SFT have been detected in Agropyron cristatum (crested wheatgrass) (Wei et al., 2000). LpFT1, LpFT4, Lp1-SST, LpVINV, LpCWINV, and LpFEH all represent single copy genes in the L. perenne genome as evidenced by single strongly hybridizing bands in Southern hybridization analyses, although this conclusion is based on relatively high stringency processing conditions (Figure 5). Weakly hybridizing bands were also observed, and may be indicative of cross-hybridization to diverged paralogous sequences such as other FTs, FEHs, and INVs.

Figure 5.

Genomic organization of fructan metabolism genes (LpFT1 (Lidgett et al., 2002), LpFT4, Lp1-SST, LpVINV (Johnson et al., 2003), LpCWINV and LpFEH) in Lolium perenne. Southern hybridization analysis of DNA from DH genotype 297 of L. perenne digested with DraI, BamHI, EcoRI, EcoRV, HindIII, and XbaI in lanes 1–6, respectively.

The LpFT1 and LpVINV genes are located in the distal upper regions of L. perenne linkage groups (LG) 7 and 6, respectively, based on restriction fragment length polymorphism (RFLP) mapping within the framework of the p150/112 reference genetic map (Lidgett et al., 2002; Johnson et al., 2003). The locations of the Lp1-SST, LpFT4, LpFEH and LpCWINV genes were determined on the parental genetic maps (Faville et al., 2004) obtained for the L. perenne genetic mapping population derived from a pair-cross between the low and high fructan content genotypes North African6 (NA6) and Aurora6 (AU6), respectively (F1(NA6 × AU6)) (Figure 6). The RFLP-detected genetic loci corresponding to LpFEH (xlpfeh) are located on LG3 of both parental maps, while the locus detected by LpCWINV (xlpcwinv) is located on LG6 of the AU6 parental map. The single nucleotide polymorphism (SNP) locus detected by Lp1-SST (xlp1-sst) is located on LG7 of the NA6 parental map, while LpFT4 (xlpft4) is located on LG3 of the AU6 parental map. The LpFT1 and LpVINV genes, previously assigned to LGs 7 and 6, respectively, in the p150/112 reference genetic map were also assigned as RFLP loci to the distal regions of LGs 7 and 6 of the NA6 parental map (Figure 6), providing confirmation of the location of these genes in a second mapping cross (Lidgett et al., 2002; Johnson et al., 2003; Faville et al., 2004). Lp1-FFT and Lp6G-FFT have been mapped within another reference mapping population derived from a cross between inbred genotypes of the cultivars Aurora and Perma which also display differing water soluble carbohydrate contents (F2(Aurora × Perma)) (Toshihiko Yamada, personal communication). The Lp1-FFT gene was assigned to the upper part of LG7, in a region that did not coincide with locations for fructan content quantitative trait loci (QTLs), while the Lp6G-FFT gene mapped to LG3 close to the location of a QTL for low DP fructan content.

Figure 6.

Genetic maps of Lolium perenne LG3 and LG6 for the NA6 and AU6 parents and LG7 of the NA6 parent based on the F1(NA6 × AU6) cross. Genomic DNA-derived SSR markers are indicated as xlpssr loci using the nomenclature described by Jones et al. (2002b). EST-RFLP markers are indicated with xlp (Lolium perenne) prefixes and gene-specific abbreviations as described in Faville et al. (2004). The xlpfeh, xlpvinv (Johnson et al., 2003), xlpcwinv, xlp1-sst, xlpft4 and xlpft1 (Lidgett et al., 2002) loci are indicated in bold.

Comparative genomics analyses of selected L. perenne sequences with putative wheat orthologues revealed that the wheat FEH gene Ta1-FEH shares 98% nucleotide sequence identity with a wheat EST (accession BE488501) that was deletion-mapped to the short arms of the homeologous group 6 chromosomes. By contrast, LpFEH appears to correspond to a second wheat EST (BE443203) mapped to the wheat homeologous group 3 chromosomes, which are the syntenic counterparts of L. perenne LG3 (Jones et al., 2002a; Van den Ende et al., 2003). This provides evidence for the existence of multiple FEH homologues in L. perenne. LpCWINV revealed highest sequence similarity to wheat ESTs (accessions BF292170 and BE606542), which map to a number of wheat chromosomes, including homeologous group 4 chromosomes, suggesting the presence of a complex gene family of LpCWINV homologues in wheat. However, the observed map locations on the homeologous group 4 chromosomes are consistent with the comparative genetic relationships with L. perenne LG4, on which LpCWINV is located.

Expression analysis of fructan metabolism genes

Expression analyses of fructan metabolism genes have shown that transcript profiles are generally consistent with both enzymatic activity measurements and levels of fructan accumulation, suggesting that these genes are primarily regulated at the transcriptional level (Lüscher et al., 2000a; Van den Ende et al., 2000; Wang et al., 2000; Koroleva et al., 2001; Lidgett et al., 2002; Van Laere and Van den Ende, 2002; Chalmers et al., 2003; Johnson et al., 2003). In L. perenne, LpFT1, Lp1-SST, and LpVINV transcripts accumulate in the base of the youngest leaf and in the sheaths of the more mature leaves, representing organs of FT activity, fructan accumulation, and remobilization (Morvan-Bertrand et al., 2001; Pavis et al., 2001a; Lidgett et al., 2002; Chalmers et al., 2003; Johnson et al., 2003). In H. vulgare 1-SST transcript and enzyme activity are both under tight regulation, responding rapidly to changes in light, while 6-SFT shows much slower regulation at both levels and a significantly longer enzyme half-life than 1-SST (Nagaraj et al., 2004).

CWINVs and VINVs require additional post-translational regulation, because these proteins are particularly stable and require rapid down-regulation in response to specific stimuli, which cannot be achieved through transcriptional repression alone (Link et al., 2004). Different INV isoforms are expressed in specific cell types and under different conditions suggesting distinct physiological roles for each isoform (Xu et al., 1996). In L. temulentum, for example, two VINVs have been isolated, which have contrasting expression profiles. LtINV1:4 shows decreased expression during fructan synthesis and enhanced expression during remobilization, while LtINV1:2 expression is predominantly in root tissues (Gallagher et al., 2004). It is difficult to correlate supporting enzymatic data with expression analysis for INVs because of the presence of multiple isoforms and the similarity in activities of INVs and FTs in vitro (Obenland et al., 1993).

Functional characterization of fructan metabolism genes

Functional characterization of plant fructan metabolism genes has been performed by enzyme assays with recombinant proteins produced in yeast, transient gene expression assays with isolated protoplasts or gain-of-function analysis through transgenesis in heterologous plant species (Sprenger et al., 1995; Koops and Jonker, 1996; Hellwege et al., 1997; Sprenger et al., 1997; Vijn et al., 1997; Hochstrasser et al., 1998; Sevenier et al., 1998; Van der Meer et al., 1998; Hellwege et al., 2000; Lüscher et al., 2000b; Cairns, 2003; Ritsema et al., 2003; Hisano et al., 2004a). The methylotrophic yeast Pichia pastoris has been widely used to generate recombinant protein for functional analysis, as synthesized protein is secreted and is consequently easily purified; P. pastoris does not secrete sucrose-metabolizing enzymes like invertases; and it has the ability to perform eukaryotic post-translational modifications including protein processing, folding, and glycosylation that would not occur in bacterial expression systems (Hochstrasser et al., 1998).

Functions for five of the eight fructan metabolism cDNAs isolated from L. perenne have been verified using the P. pastoris system. Lp1-SST utilizes sucrose to produce predominantly 1-kestose and smaller amounts of 1,1-kestose (Figure 7). LpVINV and LpCWINV are typical INVs, hydrolysing sucrose into its constituent monosaccharides, glucose and fructose (Figure 7). In contrast to an acid INV from H. vulgare (accession AJ623275), the L. perenne INVs show an additional 1-FEH-like activity, hydrolysing small inulin fructans into glucose and fructose (Figure 7) (Nagaraj et al., 2005). Activities of gene products from cDNA sequences previously annotated as a 6-FT (accession AF494041) and a 1-SST (AF492836) have been determined, through P. pastoris recombinant protein assays, to be a 1-FFT (Lp1-FFT AB186920) and 6G-FFT (Lp6G-FFT AB125218), respectively (Toshihiko Yamada, personal communication).

Figure 7.

Functional analysis of recombinant Lp1-SST (Chalmers et al., 2003), LpVINV (Johnson et al., 2003), and LpCWINV protein produced in Pichia pastoris. Recombinant protein was incubated at RT with 100 mm sucrose (S) or 1,1-kestose (1,1-K), and products of the reaction analysed by HPAEC-PAD at various time points. P. pastoris transformed with an empty vector (pPICZαA) was used as a negative control. G glucose, F fructose, 1-K 1-kestose.

The activity of this collection of enzymes may explain the production of the inulin series and inulin neoseries fructans in L. perenne, and the degradation of sucrose, the substrate of fructan biosynthesis. To date, no sequence from L. perenne has been described that can explain the production of β(2-6) linked levan neoseries fructans. The production of levan fructans is the result of 6-SFT activity in other graminaceous species. However, the absence of bifurcose in L. perenne suggests that another enzyme may be responsible for the levan neoseries fructans in this species (Sprenger et al., 1995; Wei et al., 2000; Wei and Chatterton, 2001). The FT homologues LpFT1 and LpFT4 remain potential candidates for this role.

Expression of chimeric genes encoding bacterial and plant FT enzymes in transgenic ryegrass plants has been described, exploiting established transformation methodologies in Lolium (Spangenberg et al., 1995; Ye et al., 2001; Hisano et al., 2004a). However, no report on detailed functional analysis of L. perenne fructan metabolism genes by de-regulation or loss-of-function through transgenic gene silencing approaches, or of any other grass or cereal fructan metabolism gene in the respective host plant, is yet available. To this end, hairpin RNA vectors have been generated for the targeted down-regulation of fructan metabolism genes (Lp1-SST, Lp1-FFT, Lp6G-FFT, LpFEH, LpFT1, LpFT4 and LpVINV) in L. perenne for detailed in planta functional analysis.

Promoter analysis

Current evidence suggests plant fructan metabolism genes are regulated predominantly at the transcriptional level and therefore the promoters of these genes are likely to play an important role in regulating fructan metabolism. Promoter sequences from four fructan metabolism genes of temperate grasses and cereals have been reported: Hv6-SFT, Fa1-SST, LpFT1, and Lp1-SST (Nagaraj et al., 2001; Lidgett et al., 2002; Chalmers et al., 2003; Nagaraj et al., 2003). In addition, 900 bp of promoter sequence has been isolated for the L. perenne FT homologue LpFT4.

The Hv6-SFT promoter, contains regulatory elements required to confer sucrose and light-inducible expression, including I-boxes, GT-1 binding sites, GATA boxes and TGACG motifs, which have all been shown to be functionally relevant in light-regulated promoters (Nagaraj et al., 2001). Similarly, regulatory elements conferring light-regulated, stress-inducible and phloem-specific gene expression have been identified in silico in LpFT1, LpFT4, and Lp1-SST promoter sequences, in addition to basic eukaryotic promoter elements, including CAAT and TATA boxes (Figure 8). The Lp1-SST promoter sequence contains light, stress and sucrose-related elements, while the LpFT1 promoter sequence shows predominantly light-regulated elements. Both promoter sequences include a low temperature responsive element (LTRE). The LpFT1 promoter also includes GATA motifs (A[N]3GATA) and ASL boxes (GCA[N]6-12GCA) for phloem-specific expression which have also been reported in promoters from two INV genes (Hedley et al., 2000). The LpFT4 promoter contains GT-1 core motifs and GATA boxes for light-regulated expression and stress-related elements, including MYB, MYC, and ABA responsive elements. Fructans are known to accumulate in plants upon illumination, supply of sucrose and during cold or water stress (Wagner et al., 1983; Pontis, 1989; Tognetti et al., 1990; Volaire and Lelievre, 1997). The regulatory elements present in the promoter sequences of fructan metabolism genes from temperate grasses and cereals reflect these patterns of fructan accumulation.

Figure 8.

Regulatory elements in promoter sequences of FT homologues of Lolium perenne. (a) 1.6 kb Lp1-SST promoter (b) 1.6 kb LpFT1 promoter, and (c) 0.9 kb LpFT4 promoter. Details of elements can be found at http://www.dna.affrc.go.jp/PLACE/(Higo et al., 1999).

The nature of sugar-mediated regulation of fructan metabolism gene expression has been further investigated in H. vulgare and T. aestivum. Induction of FT expression by the disaccharides sucrose and trehalose, in addition to weaker induction by the monosaccharides glucose and fructose, indicates disaccharide-mediated regulation of FTs in these two species (Muller et al., 2000; Noel and Tognetti, 2001). Sucrose sensing is hexokinase independent in H. vulgare and T. aestivum and involves protein kinase/phosphatase activities (Martinez et al., 2001; Noel and Tognetti, 2001). Nitrate has also been demonstrated as a negative signal for fructan synthesis, independent from the sugar signalling pathway (Morcuende et al., 2004).

Biotechnology applications in molecular plant breeding of ryegrasses

L. perenne is the predominant grass species in temperate pastures worldwide and is limited by seasonal changes in nutritive value and persistence. Fructans provide a readily available source of energy for grazing ruminants and high-fructan content has been associated with improved drought survival in grasses (Volaire and Lelievre, 1997). The best forage varieties yield on average 200 g/kg dry matter in the form of sucrose and fructan, while leaf tissue has the capacity to accumulate at least twice this amount (Cairns et al., 2002). Only small changes in carbon metabolism are required to achieve significant changes in fructan concentration (Smith et al., 2001, 2002), which translate into increased stocking rates, live weights, and milk production.

Substantial preexisting genetic variation for fructan content may be modified by the selection of genetic markers linked to appropriate QTLs. The large-scale development of candidate gene-associated SNP markers will permit diagnostic tests for superior allele content for target traits (Forster et al., 2004). The production of transgenic grasses through gene technology using existing methodologies (Spangenberg et al., 1995; Wang et al., 2001; Ye et al., 2001; Hisano et al., 2004a) offers the opportunity to generate unique genetic variation, when the required variation is either absent or has very low heritability.

Conclusions and future directions

Genes encoding fructan metabolism genes have been isolated from a number of temperate grass species, including L. perenne. Characterization of the sequences from L. perenne has shown they predominantly correspond to single copy genes, and in some cases map to genomic regions associated with phenotypic variation for carbohydrate content. Their transcript levels correlate with fructan metabolism enzyme activities in plant organs in which fructans typically accumulate. Their promoter sequences reveal regulatory elements associated with gene expression under conditions of fructan accumulation. Activities of the enzymes they encode are sufficient to explain the synthesis of inulin series and inulin neoseries fructans in L. perenne. The exact nature of the activity producing levan-type fructans in L. perenne remains elusive. The availability of this range of fructan metabolism genes in L. perenne opens up opportunities for the production of transgenic grasses with precisely altered endogenous fructan pools. Promoter sequences available for some of these genes will enable targeted and coordinated modification of fructan metabolism in transgenic L. perenne plants. The molecular dissection of fructan metabolism in transgenic ryegrass through targeted gene silencing of specific genes, individually and in combination, will allow us to undertake a detailed in planta functional analysis of fructan type and content, including presence-absence (e.g. Lp1-SST), DP (e.g. LpFEH and Lp1-FFT), and linkage type (e.g. Lp6G-FFT). The modification of fructan accumulation and remobilization in transgenic grasses will allow the determination of the physiological roles that fructans play under water-limitation stress and other abiotic conditions. The obtained knowledge will ultimately inform molecular breeding approaches for the development of cultivars with enhanced nutritive value, tolerance to abiotic stress and persistence.


We thank Xenie Johnson, Katherine Terdich, Noel Cogan, Anita Vecchies, Bec Ponting and Michelle Drayton for their contribution to the work reviewed. This work was supported by the Department of Primary Industries, Victoria, Australia, and the Australian Molecular Plant Breeding Cooperative Research Centre.