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Plant Physiol. Apr 2008; 146(4): 1821–1833.
PMCID: PMC2287353

The Genetics and Transcriptional Profiles of the Cellulose Synthase-Like HvCslF Gene Family in Barley1,[OA]


Cellulose synthase-like CslF genes have been implicated in the biosynthesis of (1,3;1,4)-β-d-glucans, which are major cell wall constituents in grasses and cereals. Seven CslF genes from barley (Hordeum vulgare) can be divided into two classes on the basis of intron-exon arrangements. Four of the HvCslF genes have been mapped to a single locus on barley chromosome 2H, in a region corresponding to a major quantitative trait locus for grain (1,3;1,4)-β-d-glucan content. The other HvCslF genes map to chromosomes 1H, 5H, and 7H, and in two cases the genes are close to other quantitative trait loci for grain (1,3;1,4)-β-d-glucan content. Spatial and temporal patterns of transcription of the seven genes have been defined through quantitative polymerase chain reaction. In developing barley coleoptiles HvCslF6 mRNA is most abundant. Transcript levels are maximal in 4- to 5-d coleoptiles, at a time when (1,3;1,4)-β-d-glucan content of coleoptile cell walls also reaches maximal levels. In the starchy endosperm of developing grain, HvCslF6 and HvCslF9 transcripts predominate. Two peaks of transcription are apparent. One occurs just after endosperm cellularization, 4 to 8 d after pollination, while the second occurs much later in grain development, more than 20 d after pollination. Marked varietal differences in transcription of the HvCslF genes are observed during endosperm development. Given the commercial importance of cereal (1,3;1,4)-β-d-glucans in human nutrition, in stock feed, and in malting and brewing, the observation that only two genes, HvCslF6 and HvCslF9, are transcribed at high levels in developing grain is of potential relevance for the future manipulation of grain (1,3;1,4)-β-d-glucan levels.

Plant cell walls are chemically complex, diverse structures that are modified throughout the processes of cell division, cell growth, and differentiation, and in response to abiotic and biotic stresses. Primary walls are composed mainly of cellulose and a range of noncellulosic polysaccharides, together with smaller amounts of proteins and other components (Carpita et al., 2001; Trethewey et al., 2005). Although the chemical structures of most wall polysaccharides have been determined (Fincher and Stone, 2004), there is relatively little information on the enzymes involved in their biosynthesis. Within the overall glycosyltransferase (GT) class of carbohydrate-modifying enzymes (Coutinho and Henrissat, 1999; http://afmb.cnrs-mrs.fr/CAZY/) two distinct groups of enzymes have been implicated in wall polysaccharide biosynthesis, namely the type I polysaccharide synthases and the type II GTs.

Type II GTs are membrane-bound enzymes that generally contain a single transmembrane helix near their NH2 termini. The type II enzymes were initially believed to transfer glycosyl substituents onto a polysaccharide backbone from the sugar nucleotide donor in a single catalytic event (Farrokhi et al., 2006), as exemplified by the α-d-xylosyltransferases that add single α-d-xylosyl residues to the (1,4)-β-d-glucan backbone of wall xyloglucans (Faik et al., 2002). However, it has been shown recently that GAUT1, a type II galacturonyltransferase of the GT8 family, is capable of mediating the synthesis of the homogalacturonan backbone of pectic polysaccharides (Sterling et al., 2006). Similarly, type II GTs have been implicated in the synthesis of the (1,4)-β-d-xylan backbone of heteroxylans of the wall (Lee et al., 2007; Mitchell et al., 2007; Pena et al., 2007).

The type I polysaccharide synthase group includes enzymes with an action pattern based upon the iterative transfer of glycosyl residues from sugar nucleotide donors onto main chain backbones of wall polysaccharides (Farrokhi et al., 2006). These are large integral membrane proteins, containing up to 1,000 amino acid residues and up to eight transmembrane helices, and were traditionally believed to catalyze the synthesis of the backbones of wall homopolysaccharides such as cellulose or of wall heteropolysaccharides such as heteroxylans, xyloglucans, or glucomannans (Doblin et al., 2002). The type I polysaccharide synthases are encoded by members of a large multigene family known as the cellulose synthase (CesA) superfamily (Richmond and Somerville, 2000). Within the CesA gene superfamily, subgroups can be identified through sequence alignments. These include the cellulose synthase subfamily (CesA) and Cellulose synthase-like (Csl) subfamilies A to H, each of which consists of multiple genes (Richmond and Somerville, 2000). For example, in rice (Oryza sativa) there are 37 Csl genes in total, while in Arabidopsis (Arabidopsis thaliana) there are 30 (Hazen et al., 2002; Somerville et al., 2004). Surprisingly few Csl genes have been assigned a specific function in plant cell wall biosynthesis. Members of the CslA subfamily encode (1,4)-β-d-mannan synthases (Dhugga et al., 2004; Liepman et al., 2007), and the CslC group is believed to encode an enzyme that directs the synthesis of the (1,4)-β-d-glucan backbone of xyloglucans (Cocuron et al., 2007). Using a comparative genomics approach, Burton et al. (2006) showed that a cluster of the monocot-specific CslF genes in rice is located in a genomic region corresponding to one containing a major quantitative trait locus (QTL) for grain (1,3;1,4)-β-d-glucan content in barley (Hordeum vulgare). Following expression of rice CslF genes in Arabidopsis and the detection of (1,3;1,4)-β-d-glucan in the walls of transgenic Arabidopsis lines, it was concluded that the rice CslF genes encode polysaccharide synthases that are essential for the synthesis of the (1,3;1,4)-β-d-glucans of monocot cell walls, although the participation of other enzymes or ancillary proteins could not be precluded (Burton et al., 2006).

Here, we have examined the CslF gene subfamily of barley. Seven HvCslF genes and their corresponding cDNAs have been isolated and sequenced, and the genes have been assigned positions on genetic maps from a ‘Clipper’ × ‘Sahara’ mapping population. Differential rates of transcription of individual members of the gene subfamily have been quantitated in a number of organ and tissue extracts from barley, and patterns of gene transcription have been monitored in growing coleoptiles and during endosperm development.


Cloning Barley HvCslF cDNAs and Genes

National Center for Biotechnology Information EST databases were screened for barley EST sequences with the rice full-length CslF deduced amino acid sequences using blastx and the previously isolated barley CslF sequences using blastn and blastx. Where necessary, EST sequences for wheat (Triticum aestivum) were isolated using the same procedure to extend the predicted sequences for the barley CslF cDNAs. Given that homologous genes between wheat and barley often show 98% to 99% sequence identity, it was hoped that this procedure would enable us to design PCR primers from wheat CslF ESTs where no barley equivalent was found. This approach was successful in extending the sequences of both HvCslF6 and HvCslF9. No barley or wheat ESTs corresponding to OsCslF7 were present in the public databases so degenerate oligonucleotides (ATCGCCGGSGAGCTCTGGTT and TTSCGGCAGAASGGCACCCA, sense and antisense, respectively) were designed to a region conserved between OsCslF1, OsCslF2, OsCslF4, OsCslF6, and OsCslF7 genes and used to amplify sequences from seedling cDNA. HvCslF7-specific sequences were identified and extended to a full-length cDNA using 5′ and 3′ RACE. During 5′ RACE experiments products were obtained, which upon sequencing did not match to any of the previously identified HvCslF cDNAs. Extension of this fragment using further rounds of 5′ and 3′ RACE resulted in the acquisition of the full-length cDNA encoding HvCslF10.

When the nucleotide sequences of the barley HvCslF cDNAs are aligned, sequence identity values of 53% to 69% are observed between the individual barley genes (Table I).

Table I.
Sequence similarities between individual barley HvCslF genes and between their products

The HvCslF genes corresponding to the seven cDNAs were isolated by screening bacterial artificial chromosome (BAC) libraries or by PCR amplification from genomic DNA preparations, using gene-specific primers. The BAC clone number P578K13 from ‘Morex’ was found to carry two HvCslF genes, which were subsequently designated HvCslF3 and HvCslF4, and the BAC clone number P61G5, carried the HvCslF9 gene. The complete sequences for these genes were obtained from cDNA, genomic, and PCR sequencing.

HvCslF Gene Nomenclature

The nomenclature of the barley genes is based on the nomenclature of the likely orthologs from rice, which were assigned numbers in the rice genome sequencing project. The designated, putative orthologs were selected on the basis of location on syntenous regions of rice and barley chromosomes and on sequence similarities. An unrooted phylogenetic tree (Fig. 1) indicates that barley does not have genes with close sequence similarity with the rice OsCslF1 and OsCslF2 genes. As a result, no barley genes are designated CslF1 or CslF2. However, barley genes that are apparently orthologous to the rice OsCslF3, OsCslF4, OsCslF6, OsCslF7, OsCslF8, and OsCslF9 genes were identified (Fig. 1). The rice OsCslF5 gene is believed to be a pseudogene, because it contains an internal stop codon. No barley gene obviously orthologous to the rice OsCslF5 gene was detected. On that basis the barley HvCslF5 designation has not been used. Barley has an additional gene that is designated HvCslF10; this gene is most similar to HvCslF3 (Fig. 1). Overall, the identification of the barley and rice orthologs by genome location and sequence gave consistent results.

Figure 1.
An unrooted phylogenetic tree of CslF genes from barley (HvCslF) and rice (OsCslF). Amino acid sequence identities were used to identify potential orthologs between the gene families of barley and rice. The rice gene numbers are as follows: OsCslF1, Os07g36700; ...

The sequences of the barley HvCslF genes have been deposited in the public databases, under the following accession numbers: HvCslF3, EU267179; HvCslF4, EU267180; HvCslF6, EU267181; HvCslF7, EU267182; HvCslF8, EU267183; HvCslF9, EU267184; and HvCslF10, EU267185.

Structure of HvCslF Genes

Comparisons of the nucleotide sequences of the seven HvCslF cDNAs and genes allowed the identification of introns, all of which are flanked by consensus intron processing motifs. With the exception of the HvCslF7 gene, which has a single intron, the barley HvCslF genes contain two introns toward the 5′ ends of the genes (Fig. 2). The locations of the introns are conserved both in the barley gene family and in the OsCslF gene family from rice (Fig. 2). The introns in the barley genes range in size from 138 to over 5,000 bp, while those of the rice OsCslF genes are from 91 to 4,489 bp in length (Fig. 2).

Figure 2.
Structures of the barley (yellow) and rice (green) CslF genes. The positions of introns are indicated by the triangles and the length of the introns in base pairs is indicated within each triangle. The red bars show the positions of the D,D,D,QVRRW motifs, ...

Mapping the Barley HvCslF Genes

Southern hybridization analyses of DNA preparations from wheat-barley addition lines (Islam et al., 1981) allowed the HvCslF genes to be assigned to chromosomes (data not shown). The positions of these genes within chromosome arms were subsequently mapped using RFLPs, which placed HvCslF3, HvCslF4, HvCslF8, and HvCslF10 in the centromeric region of chromosome 2H, HvCslF6 to a position near the centromere of chromosome 7H, and HvCslF7 on the long arm of chromosome 5H (Fig. 3). No RFLP between ‘Clipper’ and ‘Sahara’ was found for HvCslF9 and so the gene was mapped by single nucleotide polymorphism genotyping. Sequencing of the HvCslF9 gene from ‘Clipper’ and ‘Sahara’ resulted in the identification of a single nucleotide polymorphism in the first intron at the nucleotide 257 bp from the 5′ end of the intron and 472 bp from the ATG translation start codon. At this position, the ‘Clipper’ allele has a C and the ‘Sahara’ allele has a T. Primers designed to flank this region were 5′-CCCCACTTGATCGAACCCTTAC-3′ and 5′-GCACTGGCTACAGATCAAGATCCTAC-3′. These primers were used to amplify a 750-bp fragment of the HvCslF9 gene from each of 45 ‘Clipper’ × ‘Sahara’ doubled haploid lines that are known to carry recombination events on chromosome 1H. After sequencing and scoring the fragments from each of the 45 lines the HvCslF9 gene was assigned to the short arm of chromosome 1H, near the centromere (Fig. 3).

Figure 3.
Genetic maps of barley chromosomes 1H, 2H, 5H, and 7H showing the positions of barley HvCslF genes as mapped in a ‘Clipper’ × ‘Sahara’ population. Additional loci are shown to provide a genetic context for the map ...

No recombination was observed between the HvCslF4, HvCslF8, and HvCslF10 genes, all of which mapped to the centromeric region of chromosome 2H (Fig. 3). The HvCslF3 gene could not be mapped directly, but it must also be at this position, given that it is on the same barley BAC as HvCslF4. Barley and rice genes tentatively identified as orthologs were generally located in syntenous regions of the rice and barley genomes (Stein et al., 2007), although chromosomal translocations coupled with local breakdown of colinearity made it difficult to make unequivocal conclusions in some cases (data not shown).

Transcript Profiling in Different Tissues and Organs

When mRNA abundance in extracts from 16 tissues was examined by real-time quantitative PCR (Q-PCR) with gene-specific primers, it became apparent that the HvCslF6 gene was transcribed at relatively high levels in most of the tissues examined (Fig. 4A). This agrees with the expression level as judged by EST abundance in public databases, where HvCslF6 ESTs predominate in most libraries. Transcript levels for the HvCslF6 gene were highest in extracts of the first leaf base, but the transcripts were also detected in other vegetative tissues (Fig. 4A). Levels of HvCslF6 mRNA were generally low in preanthesis floral tissues and in mature leaves. The other HvCslF mRNAs were relatively abundant in the first leaf base, root tips, coleoptiles, peduncles, and stems, but were found at low levels in floral tissues and early developing grain (Fig. 4B). The levels of HvCslF7 transcripts were generally low, but HvCslF7 mRNA could nevertheless be detected in many of the extracts (Fig. 4C).

Figure 4.
A, Normalized levels of HvCslF6 gene transcripts (arbitrary units) in a range of tissues. These transcripts were detected in all tissues examined and very high levels of HvCslF6 mRNA were detected in some tissues. Error bars on all Q-PCR plots indicate ...

When the data were expressed as a percentage of maximal normalized levels (Burton et al., 2004), the HvCslF4, HvCslF6, HvCslF8, and the HvCslF10 mRNAs were highest in extracts from the first leaf base of young seedlings (Fig. 4D). Transcripts of the HvCslF9 gene were highest in grain, while HvCslF3 and HvCslF7 transcripts were most abundant in coleoptile and peduncle extracts, respectively (Fig. 4D). In developing barley coleoptiles, transcripts for the HvCslF6 gene were the most abundant, although HvCslF3, HvCslF8, and HvCslF9 mRNAs were also detectable (Fig. 5).

Figure 5.
Normalized expression levels for the barley HvCslF genes in developing coleoptiles at various times (days) after the initiation of germination.

Transcript Profiling in Developing Endosperm

The abundance of transcripts for individual members of the HvCslF gene family were monitored independently during the development of barley endosperm, and results were compared for two barley varieties, namely the elite malting variety ‘Sloop’ and the hulless barley ‘Himalaya’. Transcripts of the HvCslF9 gene peaked at about 8 d after pollination (DAP), and appeared to be much more abundant in the variety ‘Sloop’. Transcripts of HvCslF9 decreased quickly after 8 DAP and were present at very low levels after 16 DAP (Fig. 6A). In contrast, relatively higher levels of HvCslF6 mRNA were detectable throughout endosperm development and there was a marked increase in abundance of this mRNA late in grain development, at 16 to 24 DAP (Fig. 6B). Again, differences between the varieties ‘Sloop’ and ‘Himalaya’ were detected (Fig. 6B). Transcripts of other members of the HvCslF gene family were seen at much lower, but detectable levels during endosperm development (Fig. 6C). Because a major QTL for (1,3;1,4)-β-d-glucan content of barley grain has been reported close to the gene encoding (1,3;1,4)-β-d-glucan endohydrolase isoenzyme EI (Han et al., 1995), levels of transcripts for the corresponding gene, which is designated HvGlb1, were also monitored during endosperm development. A sharp peak in mRNA abundance for HvGlb1 was detected at 12 DAP in the variety ‘Sloop’, but lower levels were detected in the variety ‘Himalaya’ (Fig. 6D). In view of the fact that the developmental pattern shown in Figure 6D depends very much on the value at 12 DAP, the experiment was repeated with the varieties ‘Sloop’ and ‘Golden Promise’, and similar results were obtained (data not shown).

Figure 6.
A, Normalized transcript levels of the barley HvCslF9 genes in developing endosperm at various times postpollination. The two data sets are for the variety ‘Sloop’ (SLP) and ‘Himalaya’ (HIM). B, Levels of the HvCslF6 gene ...

Properties of Proteins Encoded by the HvCslF Genes

The near full-length cDNAs revealed that the HvCslF genes encode family GT2 glycosyl transferase enzymes (Coutinho et al., 2003; http://www.cazy.org/) with 810 to 947 amino acid residues. Amino acid sequence identity values range from 40% to 63% overall (Table I), although higher identity values are observed in some segments of the proteins (data not shown). The putative catalytic site residues D, D, D, QxxRW (Doblin et al., 2002) are evident (Fig. 2) but no obvious signal peptides or other common peptide motifs are present. Eight trans-membrane helices are detected, with two located toward the NH2-terminal region of the proteins and six located toward the COOH terminus (Fig. 2).

In general, the deduced amino acid sequences of the proteins align closely, with the exception of HvCslF6 and HvCslF7. A region predicted to be a loop of approximately 15 to 20 residues found in all other members of the family at about position 535 (Fig. 2) is extended in HvCslF6 to over 50 amino acid residues, through the insertion of two additional sequences on either side of the sequence found in the other six proteins. In the sequence predicted for HvCslF7, several relatively short insertions and deletions are detected throughout the protein, which is shorter overall than the other members of the family (Fig. 2).


There is emerging evidence that cellulose synthase-like CslF genes mediate the synthesis of (1,3;1,4)-β-d-glucans in the cell walls of the commelinoid monocotyledon group of land plants (Burton et al., 2006). Here, seven members of the barley HvCslF gene family have been sequenced and characterized. The genes are 2.5 to 2.8 kb in length and orthologous CslF genes from barley and rice contain either one or two introns, located in the same positions (Fig. 2).

The initial identification of the rice OsCslF genes as potential participants in (1,3;1,4)-β-d-glucan synthesis was based upon QTL analyses in barley. Molecular markers flanking a major QTL on chromosome 2H for (1,3;1,4)-β-d-glucan content in ungerminated barley grain (Han et al., 1995) were used to locate the syntenous region of the rice genome on chromosome 7, in which a cluster of six rice OsCslF genes within a 100-kb region became prime candidates for a role in (1,3;1,4)-β-d-glucan synthesis (Burton et al., 2006). We have now used gene-specific probes to screen a ‘Clipper’ × ‘Sahara’ barley doubled haploid mapping population (Karakousis et al., 2003) and have placed the individual barley HvCslF genes on a genetic map (Fig. 3). Four barley genes, namely HvCslF3, HvCslF4, HvCslF8, and HvCslF10, mapped at the same position as the RFLP marker abc468, which is in the centromeric region of chromosome 2H. This map position is syntenous with the position of the cluster of six OsCslF genes on rice chromosome 7 and falls within a chromosome region in which Han et al. (1995) detected a QTL that accounted for about 19% of the variation in grain (1,3;1,4)-β-d-glucan content in a ‘Steptoe’ × ‘Morex’ mapping population. No orthologs of the rice OsCslF1 and OsCslF2 genes, which are also found in the rice cluster on chromosome 7, were detected in barley (Fig. 1) but an additional barley gene, HvCslF10, which was most similar to HvCslF3, was located in this region of barley chromosome 2H (Fig. 3).

Examination of the nucleotide sequence identities of the barley HvCslF genes in the cluster on chromosome 2H reveal values in the range of 53% to 69% (Table I). Thus, if the genes arose by duplication of a common ancestral gene in that region of the genome, there appears to have been a high degree of sequence divergence. Similarly, the transcription patterns of the HvCslF3, HvCslF4, HvCslF8, and HvCslF10 genes are divergent (Figs. 4, B and D). Transcript levels for the four genes are relatively low during endosperm development, compared with those for HvCslF6 and HvCslF9, which may mean that the products of the genes on chromosome 2H account for less of the grain (1,3;1,4)-β-d-glucan synthesis than products of the genes on chromosomes 1H and 7H, or that HvCslF transcript levels are not always indicative of the activity of encoded enzymes or other cellular components that are required for (1,3;1,4)-β-d-glucan synthesis in the grain. It must be borne in mind that the QTLs might also be related to the positions of regulatory genes that control HvCslF expression.

The other major QTL for (1,3;1,4)-β-d-glucan content in barley grain identified by Han et al. (1995) was located on chromosome 1H and accounted for about 15% of the variation in grain (1,3;1,4)-β-d-glucan content. In the ‘Clipper’ × ‘Sahara’ population used here, the HvCslF9 gene mapped near the centromere of chromosome 1H (Fig. 3). In the same region of barley chromosome 1H, Han et al. (1995) detected overlapping QTLs for (1,3;1,4)-β-d-glucan content in ungerminated barley grain (Fig. 3), for malt (1,3;1,4)-β-d-glucan content, and for malt (1,3;1,4)-β-d-glucan endohydrolase activity. They suggested that all of these QTLs may be attributable to HvGlb1, which encodes barley (1,3;1,4)-β-d-glucan endohydrolase isoenzyme EI (Woodward and Fincher, 1982; Slakeski et al., 1990). An alternative explanation, however, is that grain (1,3;1,4)-β-d-glucan content is influenced by the expression of HvCslF9 during grain development, while malt (1,3;1,4)-β-d-glucan content and malt (1,3;1,4)-β-d-glucanase activity are influenced by expression of the HvGlb1 gene during germination. Consistent with this, the test statistics (log of the odds scores) reported by Han et al. (1995) for grain (1,3;1,4)-β-d-glucan content are statistically significant across a broad region that encompasses both the HvCslF9 and HvGlb1 loci (Fig. 3), and the QTL peak for (1,3;1,4)-β-d-glucan content in grain does not coincide with the peaks for (1,3;1,4)-β-d-glucan content in malt and (1,3;1,4)-β-d-glucanase activity in malt (Han et al., 1995). Nevertheless, it is possible that (1,3;1,4)-β-d-glucan content in grain is influenced by the expression of both HvCslF9 and HvGlb1 during grain development.

If this were the case, one would expect to see transcripts for both genes in the developing endosperm. The data presented in Figure 6, A and D, confirm that both genes are transcribed in developing endosperm, albeit transiently and at different stages of endosperm development. While we have not tested for (1,3;1,4)-β-d-glucanase activity in these extracts, there is some evidence from other sources that hydrolytic enzymes might participate in polysaccharide biosynthesis during wall assembly. Thus, mutations in the Arabidopsis KOR gene, which encodes a (1,4)-β-d-glucanase, lead to large reductions in wall cellulose. The possible point of participation of hydrolytic enzymes in cellulose synthesis is unknown, but the hydrolase could be involved in trimming or editing nascent cellulose chains (Szyjanowicz et al., 2004), or possibly in releasing newly synthesized chains from the biosynthetic enzymes (Farrokhi et al., 2006).

The HvCslF6 gene mapped near the centromere of chromosome 7H (Fig. 3) in a position that corresponds well with those of previously reported QTLs affecting grain (1,3;1,4)-β-d-glucan content in a ‘Derkado’ × ‘B83-12/21/5’ mapping population (Igartua et al., 2002) and in a ‘Beka’ × ‘Logan’ population (Molina-Cano et al., 2007). A contribution by the HvCslF6 gene to these QTLs is consistent with the relatively high levels of HvCslF6 gene transcripts in the developing endosperm (Fig. 6B). Kim et al. (2004) have also reported QTLs for grain (1,3;1,4)-β-d-glucan content on chromosome 7H, one of which is close to the HvGlb2 gene that encodes (1,3;1,4)-β-d-glucanase isoenzyme EII (Fig. 3). Overall, there is considerable correspondence between published QTL map positions and the map positions of the HvCslF genes (Fig. 3).

The levels of (1,3;1,4)-β-d-glucan in cell walls of different tissues and organs of barley vary from about 5% by weight in walls from stems to as much as 70% or more by weight in walls of the starchy endosperm (Fincher and Stone, 2004). More detailed analyses of transcript abundance during the growth and development of barley coleoptiles showed that HvCslF6 mRNA was again the most abundant (Fig. 5), although significant levels of HvCslF8 and HvCslF3 transcripts were also detected (Fig. 5). The peak in HvCslF6 transcript abundance at 4 to 5 d after germination corresponds to the time when elongation of the coleoptile ceases, the leaf breaks out of the coleoptile, and maturation of the coleoptile begins (Gibeaut et al., 2005). It also corresponds to the time at which levels of (1,3;1,4)-β-d-glucan peak in coleoptiles, at about 10 mol % of the walls (Gibeaut et al., 2005).

As mentioned above, the most abundant mRNA transcripts in developing barley endosperm are those from the HvCslF6 and HvCslF9 genes (Fig. 6, A and B). However, transcriptional activities of the two genes vary both with respect to the amount of mRNA and to the timing of transcription. The transcription of the HvCslF9 gene peaks at about 8 DAP, at a stage when cellularization of the endosperm is complete and starch deposition has commenced; (1,3;1,4)-β-d-glucan is detectable in endosperm walls from 5 DAP onward (Wilson et al., 2006). In contrast, HvCslF6 transcripts are detected at much higher levels throughout endosperm development, even before (1,3;1,4)-β-d-glucans appear in the walls (Fig. 6B; Wilson et al., 2006). From about 12 to 20 DAP there are further increases in the abundance of HvCslF6 transcripts (Fig. 6B). The latter transcriptional patterns are consistent with earlier work on the development of (1,3;1,4)-β-d-glucan in barley grain, where a late increase in (1,3;1,4)-β-d-glucan was reported (Coles, 1979). Perhaps not surprisingly, variations in transcript levels of the HvCslF6, HvCslF9, and HvCslF8 genes were also observed between different barley varieties (Fig. 6, A, B, and C, respectively), consistent with the fact that QTLs for (1,3;1,4)-β-d-glucan content have been detected in chromosomal regions where these genes are located. Transcript levels of the HvCslF9 gene are much higher in the malting quality variety ‘Sloop’ than in the hulless variety ‘Himalaya’ during the cellularization stage of initial wall development (Fig. 6A), while levels of the most abundant mRNA, from the HvCslF6 gene, are higher in ‘Himalaya’ than in ‘Sloop’ during most of endosperm development, and particularly at 20 DAP (Fig. 6B). These HvCslF6 transcript levels are consistent with the generally higher levels of (1,3;1,4)-β-d-glucan in ‘Himalaya’, which are typically 6% by weight, than in ‘Sloop’, where 3% to 4% by weight (1,3;1,4)-β-d-glucan is normally found in mature grain (A.J. Box, R.A. Burton, H.M. Collins, S.A. Jobling, unpublished data).

In making these comparisons, which suggest that HvCslF transcript abundance is related temporally and spatially to the deposition of (1,3;1,4)-β-d-glucan in a range of organs and tissues, it is important to emphasize again that abundance of transcripts encoding a particular isoenzyme might not necessarily be a good indicator of enzyme activity or of (1,3;1,4)-β-d-glucan deposition in the wall. For example, the HvCslF enzymes encoded by low abundance mRNAs might be just as important as, or even more important than, the HvCslF6 and HvCslF9 enzymes for the deposition of (1,3;1,4)-β-d-glucan in developing grain. This possibility is exemplified by the observation that one starch branching isoenzyme (SBEII) makes up only 2% of the total SBE enzymes in potato (Solanum tuberosum) tubers, but has a major effect on amylose content (Jobling et al., 1999). It has also been reported that the (1,3;1,4)-β-d-glucan content of barley grain is affected in large degree by environmental factors, in particular by dry conditions toward the end of grain maturation (Fastnaught et al., 1996; Hang et al., 2007). In contrast to the situation with barley HvCesA genes, which appear to be cotranscribed to form enzyme complexes that might consist of three different HvCesA isoenzymes (Burton et al., 2004), at this stage we have not been able to detect any correlations in the transcript levels of different HvCslF genes.

Although the rice OsCslF genes appear to be involved in (1,3;1,4)-β-d-glucan synthesis (Burton et al., 2006) and the QTL analyses performed here (Fig. 3) support a similar role for the HvCslF genes in barley, it remains possible or even likely that the biosynthesis of (1,3;1,4)-β-d-glucan requires a multienzyme complex and that an individual CslF isoenzyme represents just one component of such a complex. Furthermore, there are a number of fine structural features of cereal (1,3;1,4)-β-d-glucans that have not yet been characterized at the biosynthetic level. In particular, the (1,3;1,4)-β-d-glucans of the commelinoid monocotyledons consist of a backbone of (1,3)- and (1,4)-linked β-d-glucosyl residues that are not arranged at random, but nevertheless have a degree of structural order (Staudte et al., 1983). The (1,3;1,4)-β-d-glucans from barley usually have two or three adjacent (1,4)-linked β-d-glucosyl residues separated by a single (1,3)-linked β-d-glucosyl residue. The molecules have therefore been likened to a polymer of cellotriosyl and cellotetraosyl residues linked by single (1,3)-β-linkages. The irregular distribution of the (1,3)-β-linkages along the polysaccharide chain (Staudte et al., 1983) means that there is no strictly repeating structure (Fincher and Stone, 2004). In addition, a small proportion of much longer blocks of adjacent (1,4)-linked β-d-glucosyl residues are found in the polysaccharide. The complexity of this structure can be related to the function of (1,3;1,4)-β-d-glucans in cell walls (Fincher and Stone, 2004), and is also suggestive of a similar complexity in the enzymology of (1,3;1,4)-β-d-glucan biosynthesis (Buckeridge et al., 2001), where one might again infer that multiple enzymes are required.

To further confirm the participation of HvCslF genes in (1,3;1,4)-β-d-glucan synthesis, we are attempting to manipulate levels of (1,3;1,4)-β-d-glucan, in both vegetative tissues and grain, through transgenic approaches. Altering the levels of (1,3;1,4)-β-d-glucan in walls of cereals and grasses could find applications in human and animal nutrition, or in the malting and brewing industries. Barley (1,3;1,4)-β-d-glucans are beneficial to human health, where they represent soluble dietary fiber and appear to reduce the risks of colorectal cancer, high serum cholesterol, and cardiovascular disease, obesity, and non-insulin-dependent diabetes (Brennan and Cleary, 2005). On the other hand, (1,3;1,4)-β-d-glucans are considered to be antinutritive in feed formulations for monogastric animals and have undesirable effects in cereal processing applications such as malting and brewing (Brennan and Cleary, 2005). The availability of information on the HvCslF gene family, together with the transcriptional profiles presented here, have allowed the identification of target genes for manipulation, depending on whether the objective is to increase or decrease (1,3;1,4)-β-d-glucan levels in grains, or in vegetative tissues. It should now be possible to exploit this information in breeding programs, either through a transgenic approach, or through the analysis of natural variation in HvCslF gene structure, HvCslF gene transcription rates, and (1,3;1,4)-β-d-glucan levels in mapping and mutant populations, or in germplasm collections.


Plant Material

Barley (Hordeum vulgare) ‘Golden Promise’, ‘Himalaya’, and ‘Sloop’ were grown under standard glasshouse conditions as described in Burton et al. (2004). ‘Golden Promise’ is a malting cultivar adapted to the northern United Kingdom, ‘Himalaya’ is a hulless variety that has been used extensively in studies of grain germination, and ‘Sloop’ is an elite Australian malting variety. The tissue series cDNAs used for Q-PCR were prepared as described by Burton et al. (2004). The RNA extracts from coleoptiles were prepared from young seedlings grown in vermiculite in the dark at room temperature (Gibeaut et al., 2005). Coleoptiles were dissected away from seedling leaves 1 to 7 d after imbibition of the grain. To collect endosperm from developing grain, spikes were hand pollinated and bagged at anthesis, and the developing caryopses collected at regular intervals after pollination. Conditions were maintained as described previously to ensure that the transcript analyses could be related to specific stages of grain development, as defined by high-resolution electron microscopy (Wilson et al., 2006). The entire caryopsis was collected at anthesis for the 0-DAP sample. The liquid endosperm was squeezed out of the developing caryopses up to 6 DAP, after excising the embryo with a scalpel blade. The endosperm was collected with a fine pipette tip and frozen in liquid nitrogen. The cellularized endosperm was collected from 6 DAP onward after removing the maternal pericarp and the embryo. Multiple endosperms from grain on at least three different spikes were collected and combined before RNA extraction.

RNA Isolation and cDNA Synthesis

Total RNA was extracted from all tissue homogenates using a phenol-guanidine reagent, treated with the DNA-free kit (Ambion), and used as the template for cDNA synthesis as described in Burton et al. (2004) except that Superscript III reverse transcriptase (Invitrogen) was used in preference to Superscript II and the reaction was incubated at 50°C.

SMART RACE PCR, cDNA Cloning, and Sequencing

Total RNA extracted from various tissues as described above was used as the template to make both 3′ and 5′ SMART cDNA populations using a SMART RACE kit according to the manufacturer's instructions (Clontech). Aliquots of the cDNA were used in PCR reactions with a universal primer (Clontech), gene-specific primers, and the BD Advantage2 Taq polymerase, according to the manufacturer's guidelines, to amplify fragments from the 3′ or 5′ ends of the cDNA. Products were either purified using Macherey Nagel Nucleospin columns and sequenced on an Applied Biosystems 3700 at the Australian Genome Research Facility, or cloned into pGEM-TEasy (Promega). Plasmids containing inserts were prepared using a plasmid kit (QIAGEN) according to the manufacturer's instructions and sequenced. SMART cDNA populations synthesized from RNAs extracted from the various tissues were also used as templates for the eventual isolation of full-length cDNAs, using gene-specific primers designed to the 5′ and 3′ untranslated regions.

Contig Assembly and Bioinformatics

EST sequences, RACE fragments, and genomic sequences were assembled into contigs (contiguous sequences) using the program ContigExpress, which is part of the Vector NTI Advance 9.1.0 (Invitrogen) suite of programs.

Pairwise sequence alignments were performed using the EMBOSS Pairwise Alignment Algorithm at the EMBL-EBI Web site (http://www.ebi.ac.uk/emboss/align/) with the EMBOSS::needle (global) method and default settings selected. Full-length amino acid sequences of all barley and rice (Oryza sativa) CslF proteins were aligned and a phylogenetic tree generated using the program ClustalX (Thompson et al., 1997). The phylogenetic tree was visualized using Treeview 1.6.6 (Page, 1996). Transmembrane helices were predicted using the program TMHMM (http://www.cbs.dtu.dk/services/TMHMM-2.0/; Krogh et al., 2001).

BAC Sequencing and Genomic Cloning

Three BACs carrying inserts of the barley genomic DNA from ‘Morex’ were identified after hybridization of the BAC library with a consensus barley HvCslF probe, using standard BAC library screening methods. The BAC clones were sequenced to 3× coverage and the sequences were aligned by the Beijing Genomics Institute, China. Contiguous sequences were assembled using Contig Express in the VectorNTI suite of programs (Invitrogen). Full-length genomic clones of HvCsl3, HvCsl4, HvCsl6, HvCsl7, HvCsl8, HvCsl9, and HvCsl10 were amplified from genomic DNA and sequenced on an Applied Biosystems 3700 sequencer at the Australian Genome Resource Facility, Brisbane.

Genetic Mapping

All the HvCslF genes were assigned to chromosomes using wheat-barley addition lines (Islam et al., 1981). Filters of digested genomic DNA for Southern hybridization analyses of the barley doubled haploid mapping population ‘Clipper’ × ‘Sahara’ (Karakousis et al., 2003) were hybridized with probes derived from either the promoter or the 3′ untranslated regions of each HvCslF gene. This revealed polymorphisms between ‘Clipper’ and ‘Sahara’ for the HvCslF4, HvCsl6, HvCsl7, HvCsl8, and HvCsl10 genes. All genes were mapped using Map Manager QT version 0.30 (Manly et al., 2001).

Real-Time Q-PCR

Real-time Q-PCR was carried out essentially as outlined in Burton et al. (2004) with the following modifications. To provide a template for the standard curve, between four and six 20-μL PCR reaction mixtures were combined for purification by HPLC using a HELIX DNA DVB 50- × 3.0-mm monolithic polymer reversed-phase column (Varian). Chromatography was performed using buffer A (100 mm triethylammonium acetate [Applied Biosystems] and 0.1 mm EDTA) and buffer B (100 mm triethylammonium acetate, 0.1 mm EDTA and 75% acetonitrile). The gradient was as follows: time 0 min, 10% buffer B; time 6 min, 21.5% buffer B; time 7 min, 21.5% buffer B; time 8 min, 10% buffer B; time 12 min, 10% buffer B. The flow rate was 0.45 mL/min and the temperature was 50°C. Three replicates of each of the seven standard concentrations were included with every Q-PCR experiment together with a minimum of three no-template controls. Q-PCR experiments were assembled by the liquid-handling robot CAS-1200 robot (Corbett Robotics). Three replicate PCRs for each of the cDNAs were included in every run containing: 2 μL of cDNA solution, the diluted standard, or water was used in a reaction containing 5 μL of IQ SYBR Green PCR reagent (Bio-rad Laboratories), 1.2 μL of each of the forward and reverse primers at 4 μm, 0.3 μL of 10× SYBR Green in water, and 0.3 μL of water. The total volume of the PCR reactions was 10 μL. Reactions were performed in an RG 6000 Rotor-Gene real-time thermal cycler (Corbett Research): 3 min at 95°C followed by 45 cycles of 1 s at 95°C, 1 s at 55°C, 30 s at 72°C, and 15 s at the optimal acquisition temperature described in Table II. The transcript levels of genes encoding glyceraldehyde 3-phosphate dehydrogenase, heat shock protein 70, cyclophilin, and α-tubulin were used as controls (Table II), and these correspond to those listed in table IV of Burton et al. (2004). Normalization was carried out using multiple control genes as described by Burton et al. (2004) and the final concentrations of mRNAs of the genes of interest are expressed as arbitrary units that represent the numbers of copies per microliter of cDNA, normalized against the geometric means of the three control genes that vary the least with respect to each other (Vandesompele et al., 2002).

Table II.
Gene-specific QPCR primers, with PCR product sizes in base pairs and optimal acquisition temperatures for the genes analyzed

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers EU267179 to EU267185.


We are grateful to Jacinda Rethus, Anne Medhurst, and Robin Chapple for their assistance in various aspects of the work, to Ursula Langridge for her expertise in plant care, and to Margie Pallotta for her ongoing assistance with the genetic mapping of genes.


1This work was supported by the Australian Research Council, the Grains Research and Development Corporation, the Commonwealth Scientific and Industrial Research Organization Food Futures Flagship program, and the Commonwealth Scientific and Industrial Research Organization Flagship Collaboration Fund.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Geoffrey B. Fincher (ua.ude.edialeda@rehcnif.ffoeg).

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