JXB Advance Access originally published online on September 25, 2003
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Journal of Experimental Botany, Vol. 54, No. 392, pp. 2467-2477,
November 1, 2003
© 2003 Oxford University Press
Two 
-L-arabinofuranosidase genes in Arabidopsis thaliana are differentially expressed during vegetative growth and flower development*
Received 3 March 2003; Accepted 17 July 2003
Department of Genetics, University of Melbourne, Victoria 3010, Australia
* The cDNA nucleotide sequences reported in this paper have been submitted to the GenBankTM/EMBL Nucleotide Sequence database with accession numbers AY243509 (AtASD1) and AY243510 (AtASD2). To whom correspondence should be addressed. Fax: +61 3 8344 5139. E-mail: ccobbett{at}unimelb.edu.au
Abbreviations: CAZY, carbohydrate-active enzyme; RT-PCR, reverse transcriptase polymerase chain reaction.
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Abstract |
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Glycosyl hydrolases are important mediators of plant cell wall modification during plant development. These enzymes catalyse the hydrolytic release of specific sugars, such as L-arabinose, from the polysaccharide-rich cell wall matrix. The cloning and expression analysis of two genes, AtASD1 and AtASD2, encoding putative
-L-arabinofuranosidases in Arabidopsis thaliana are reported here. AtASD1 and AtASD2 identities were assigned on the basis of homology to plant and microbial family 51 glycoside hydrolases. Using RT-PCR, RNA gel blot analysis and reporter gene expression analysis, AtASD1 and AtASD2 were shown to have different developmental expression profiles. High levels of AtASD1 promoter activity are present in multiple tissues during vegetative and reproductive growth. AtASD1 expression is particularly intense in zones of cell proliferation, the vascular system, developing and regressing floral tissues, and floral abscission zones. By comparison, AtASD2 expression is limited to the vasculature of older root tissue and to some floral organs and floral abscission zones. Key words: Abscission, Arabidopsis, arabinose, expression, gene structure, glycoside hydrolases, GUS (ß-glucuronidase), vegetative growth.
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Introduction |
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The monosaccharide L-arabinose is an integral component of various structural polysaccharide elements and glycoproteins in the plant cell wall. In higher plants, L-arabinose is found in glycan side-chains of both rhamnogalacturonan I (RG-I) and rhamnogalacturonan II (RG-II), the two highly-branched acidic polymer species that cross-link and reinforce homogalacturonan (HG) in the pectic polysaccharide complex (Ridley et al., 2001; Willats et al., 2001b; Glushka et al., 2003). In monocot species, the predominant structural hemicellulose is glucuronoarabinoxylan (Carpita and Gibeaut, 1993), which consists of a backbone of (1->4)-ß-D-linked xylopyranosyl units that may be singly or doubly substituted with 2-O- and 3-O-linked
-L-arabinofuranosyl residues. L-arabinose also forms a significant component of arabinogalactan proteins (AGPs), as arabinogalactan side-chains covalently linked to core protein (Showalter, 2001). Collectively, L-arabinose is represented in a heterogeneous and functionally diverse group of cell wall molecules.
A fundamental characteristic of the primary cell wall is its dynamic nature. Structurally, the wall matrix comprises load-bearing cellulose microfibrils hydrogen-bonded to hemicelluloses, supported within a domain of pectic polysaccharides and glycoproteins (reviewed in Carpita and Gibeaut, 1993). This framework defines cellular form and wall mechanical strength. It also allows for a plastic environment that accommodates plant developmental change. Current models suggest that wall dynamics occur primarily through hydrolysis, glycan integration and restructuring transglycosylation within the cellulose–xyloglucan complex (Nicol et al., 1998; Thompson et al., 1997; Pauly et al., 1999; Whitney et al., 1999; Thompson and Fry, 2001). Pectin remodelling is also evident during development (Willats et al., 2001b). Interestingly, pectic (1->5)--arabinan side-chains can be developmentally regulated (Willats et al., 1999; Bush et al., 2001; McCartney and Knox, 2002). Within plants, pectic (1->5)--arabinan is also shown to have interesting patterns of distribution that coincide with cell type (Willats et al., 1998, 2001a; Ermel et al., 2000; McCartney and Knox, 2002) or particular structural aspects of individual cell surfaces (Orfila and Knox, 2000; Orfila et al., 2001). At present, little is known about the specific distribution of RG-II-associated arabinose-containing glycans. Although particular physiological roles have not yet been identified, the differential localization of arabinose-containing glycans within and between species suggests important and diverse functions.
Glycosyl hydrolases, responsible for a substantial part of sugar turnover in the cell wall, are important when considering the mechanisms of cell wall dynamics. Recently, the first genes encoding plant arabinoxylan arabinofuranohydrolase proteins, AXAH-I and AXAH-II, were described in barley (Horduem vulgare L.) (Lee et al., 2001). Preparations of AXAH-I and AXAH-II from two independent studies were shown to hydrolyse (1->2), (1->3)- and (1->5)-linked arabinofuranosyl substituents from various substrates, including p-nitrophenyl -L-arabinofuranoside, barley and wheat flour arabinoxylan, sugar beet arabinan, larch wood arabinogalactan and (1->5)-linked -L-oligoarabinosides (Ferré et al., 2000; Lee et al., 2001). In this study two cDNA sequences were identified that encode putative -L-arabinofuranosidase proteins in Arabidopsis thaliana, denoted AtASD1 and AtASD2. These sequences are highly homologous to AXAH-I and AXAH-II in barley. In this paper, gene structures and the spatial and developmental expression of AtASD1 and AtASD2 are reported, as determined by RT-PCR and promoter::GUS reporter gene expression.
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Materials and methods |
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Plant material
A. thaliana (ecotype Columbia) seeds were germinated and grown on nutrient agar medium at 20 °C under a light (150–250 µE m–2 s–1)/dark cycle of 16/8 h. For the growth of adult plants, 7 d seedlings were transferred to soil medium and incubated in continuous fluorescent illumination (230–250 µE m–2 s–1) at 21 °C.
Isolation of cDNA clones
A 
Zap II cDNA library (Arabidopsis Biological Resource Center, Columbus, OH, USA) made of mRNA from 3-d-old Arabidopsis thaliana (Colombia) seedling hypocotyls (Kieber et al., 1993) was screened. Nitrocellulose lifts of plated library dilutions were performed and hybridized with [-32P]-dATP-labelled DNA from an expressed sequence tag (97D13T7) corresponding to At3g10740. Positive clones were detected by autoradiography, purified and excised into pBluescript SK+ form for restriction digestion analyses and sequencing. An AtASD2 cDNA was amplified with polymerase chain reaction (PCR) from reverse-transcribed flower RNA. Gene-specific primers 5'-AAAATCTAGATGATTATGGACATGGAAA CATCTT-3' and 5'-AAAAGAGCTCAT GATGTTTCTCTTCACA TTG TTG -3' which contained 5' Xba1 and Sac1 restriction sites, respectively (underlined), amplified a coding sequence which included the predicted start ATG and stop codon (–5 to +2043). This fragment was ligated into the Xba1 and Sac1 sites in pBluescript SK+ (Stratagene).
Nucleotide sequencing and sequence analysis
DNA sequencing was performed with the ABI Prism BigDye terminator kit (Applied Biosystems, Foster city, CA, USA), using the dideoxyribonucleotide chain termination methodology (Sanger et al., 1977). Reaction products were purified and sent to the Australian Genome Research Facility (AGRF) for gel separation and sequence determination. Sequence data files were compiled with the SequencherTM version 3.1 (Gene Codes Corporation ©, Ann Arbor, MI, USA) program. Genomic nucleotide sequences were obtained with BLASTN searches of GenBank through the National Center for Biotechnology Information web server (NCBI; http://www.ncbi.nlm.nih.gov). Family 51 glycoside hydrolase protein sequences were obtained from GenBank and the Carbohydrate Active Enzyme database (Coutinho and Henrissat, 1999; URL: http://afmb.cnrs-mrs.fr/CAZY/).
Sequence comparisons and phylogenetic tree
An alignment of the deduced AtASD1 and AtASD2 protein sequences with barley AXAH-I and AXAH-II was generated using the Pileup program through WebANGIS (http://www.angis.org.au/WebANGIS) and schematically modified using MacBoxshade version 2.15. (Kay Hoffman: Boxshade; Michael D Baron: MacBoxshade derivative). Multiple sequence alignments for phylogenetic analysis were performed using the ClustalW alignment tool in MacVector 7.0TM software (© Oxford Molecular group, 2000). An unrooted phylogenetic tree was constructed with the PAUP (Swofford, 1998) software, using maximum parsimony methodology. Bootstrap values were generated (repetitions=1000).
Reverse transcriptase-polymerase chain reaction (RT-PCR) analysis
First strand complementary DNA was synthesized from total RNA samples (5 µg) using the SuperScriptTM First-Strand Synthesis System for RT-PCR system (Invitrogen, Carslbad, CA, USA). Subsequent PCR amplification used Taq DNA polymerase. A 674 bp AtASD1 transcription product was detected using primers 5'-CATCGGGAGGTGTCGGAGTTTAT-3' and 5'-GGCATCTCCTC GGGCAAACTCG-3', whilst AtASD2 primers 5'-AAGACGCCAT TGTAACACTGC-3' and 5'-ATAAGGTCTTCCTGCTCAGTC-3' amplified a 1.8 kb product. A 200 bp ACT2 product was amplified using primers 5'-GGTAACATTGTGCTCAGTGGTGG-3' and 5'-CTCGGCCTTGGAGATCCACATC-3', as an internal control. Each primer pair was designed to span introns within the genomic sequence of each gene. Control genomic products of 1.4 kb, 3.3 kb and 0.3 kb were expected for AtASD1, AtASD2 and ACT2, respectively. PCR amplifications were performed for 30 cycles (AtASD1 and ACT2) and 35 cycles (AtASD2), using 1 µl template at appropriate dilutions and an annealing temperature of 57 °C.
Promoter–GUS gene fusion constructs
The forward and reverse primers 5'-AAAAGTCGACCCAATG ACCTACCAGAAC-3' and 5'-AAAAGGATCCCACCACATCCA AATAAACC-3', and forward and reverse primers 5'-AAAA GTCGACGGTCGGAAAAAGTCAGATGT-3' and 5'-AAAA GGATCCAATCACCAAATTTAACCAGAGAA-3', were used to amplify upstream regions of the AtASD1 (–2854 to –3) and AtASD2 (–2016 to –1) genes, respectively. Primer-incorporated restriction sites SalI and BamHI were used to clone respective fragments into pBluescript for DNA sequence confirmation and ligation between the SalI and BamHI sites in pBI101 (Clontech, Palo Alto, CA, USA), in fusion with the GUS::NOS terminator cassette. The resulting constructs were transformed into wild-type Col-0 plants via the Agrobacterium-mediated floral dip transformation method (Clough and Bent, 1998). Kanamycin-resistant primary transformants were self-fertilized for selection of homozygous transgenic lines.
GUS histochemistry
For histochemical localization of GUS activity, tissues were incubated in GUS staining buffer [0.2% Triton-X 100 (v/v), 50 mM NaPO4 at pH 7.2, 2 mM potassium ferrocyanide, 2 mM potassium ferricyanide, 2 mM X-Gluc in dimethyl formamide] at 37 °C for 12 h. Prior to incubation, tissues were subject to 30 s of vacuum to facilitate penetration of the substrate. Chlorophyll was removed from samples by incubation in 70% ethanol. Tissues were photographed either suspended in ethanol or mounted in DPX (BDH Chemicals, Poole, UK) on glass slides.
Northern analysis
To test AtASD1 and AtASD2 gene expression responses to plant hormones, 7 d seedlings were incubated for 24 h with each of the following; indole acetic acid, 2,4-dichlorophenoxyacetic acid, gibberellic acid, abscisic acid, kinetin, and epibrassinolide at concentrations of 0.1 µM, 1.0 µM and 10.0 µM. Hormone treatments had previously been confirmed to elicit physiological responses, through seed germination and seedling growth studies on hormone-supplemented minimal media. Growth responses were observed at lowest concentrations of 0.1 µM and 1.0 µM (IAA only) and greater.
Total RNA was extracted from freshly harvested tissue using the RNAqueousTM (Ambion Inc. Austin, TX, USA) extraction kit and stored at –70 °C. For northern analysis, RNA fractionation was performed according to agarose-formaldehyde gel electrophoresis methodology (Ausubel et al., 1994). RNA gel blotting was performed to nylon membrane (Hybond N, Amersham). DNA probe template was radiolabelled with [32P]-dATP using the StripEZTM DNA (Ambion) labelling kit, denatured and incubated with blots in ULTRAhybTM (Ambion) hybridization buffer, as per the manufacturer’s instructions. Final washes [0.1x SSC and 0.1% SDS] were performed at 42 °C for 2x5 min intervals. AtASD1 and AtASD2 transcripts were probed with 880 bp and 800 bp DNA fragments from respective cDNA clones. Radioactivity in blots was detected upon exposure to autoradiograph film.
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Results and discussion |
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Isolation and characterization of AtASD1 and AtASD2 cDNA clones
The Arabidopsis genome contains two putative
-L-arabinofuranosidase genes designated as At3g10740 (T7M13.18) and At5g26120 (TIN24.13), which are denoted AtASD1 and AtASD2 respectively, in this report. An EST sequence (97D13T7), which corresponded to At3g10740, was used to probe an Arabidopsis hypocotyl cDNA library from which a 2.3 kb full-length cDNA clone was isolated. The nucleotide sequence of this clone (GenBank Accession No. AY243509
[GenBank]
) contains a single open-reading frame of 2073 bp. Alignment with the genomic sequence identified 17 exons, interspersed by small introns of 80–120 bp in length (Fig. 1A). An additional 451 bp intron is situated between positions –569 to –28 relative to the start ATG codon. The deduced protein sequence of 678 amino acids includes a 20 amino acid N-terminal secretion signal with a cleavage sequence LLG|SC, as predicted by PSORT. Four N-glycosylation site consensus sequences, N-X-(T/S), are located at positions 181, 362, 523, and 555 of the unprocessed peptide.
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The cDNA of AtASD2 was amplified using RT-PCR with primers encompassing the start and stop codons predicted from the genomic DNA sequence. A 2.0 kb cDNA product of AtASD2 was isolated and sequenced (GenBank Accession No. AY243510 [GenBank] ). The gene structures of AtASD1 and AtASD2 are conserved, both in the number and relative positions of the introns (Fig. 1B). An additional 5' untranslated sequence was obtained from an independent full-length cDNA sequence (Accession No. AY063925 [GenBank] ). This allowed the identification of an intron (–520 to –27) upstream of the predicted AtASD2 start ATG (Fig. 1B), similar to that found in AtASD1. The deduced AtASD2 translation product is 674 amino acids in length and 78.3% identical to AtASD1. PSORT predicts AtASD2 has a 24-residue N-terminal signal peptide, cleaved at the sequence (SFS|VY). Seven N-glycosylation consensus motifs are present in AtASD2, three of which are conserved between the two homologues. AtASD1 and AtASD2 exhibit between 68% and 71% amino acid identity with the barley proteins, AXAH-I and AXAH-2 (Fig. 2).
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The similarity between AtASD1, AtASD2 and barley AXAH-I and AXAH-II, suggests these hydrolases are likely to have similar biochemical substrate specificities. AtASD1 and AtASD2 activities have not been directly demonstrated however, despite heterologous expression in Escherichia coli, Saccharomyces cerevisiae and Pichia pastoris (data not shown). Glycosylation may be an important post-translational modification for these proteins. Many plant glycosyl hydrolases described in the literature contain potential N-glycosylation sites (Nicol et al., 1998; Lee et al., 2001, 2003; Sampedro et al., 2001; de la Torre et al., 2002; Goujon et al., 2003) or, like AXAH-I and AXAH-II, have been shown to be glycosylated (Ferré et al., 2000; Cairns et al., 2000). The incorrect or absence of glycosylation in heterologously-expressed proteins is a likely reason for lack of detectable activity.
AtASD1 and AtASD2 have been classified as family 51 glycoside hydrolases (GH51) (Lee et al., 2001). Currently, 35 microbial and 6 plant enzymes are listed in this family, which are predominantly described as arabinosidases and -L-arabinosidases (Coutinho and Henrissat, 1999; URL: http://afmb.cnrs-mrs.fr/CAZY/). In addition, a complete, related protein sequence from tomato has been identified in Genbank (AF42998). In an unrooted phylogenetic tree these enzymes form a clustered subfamily, with bacterial and fungal enzymes forming two additional branches (Fig. 3).
In many glycoside hydrolase families, including GH51, the catalytic domain is identified by two signature glutamate residues in conserved domains (Henrissat et al., 1995). The likely catalytic acid residues in AtASD1 and AtASD2 are Glu-388 and Glu-387, respectively, while corresponding nucleophile residues are predicted as Glu-465 and Glu-464, respectively (Fig. 2). These residues were assigned on the basis of protein sequence alignments and descriptions in the literature (Henrissat et al., 1995; Zverlov et al., 1998; Shallom et al., 2002). Corresponding glutamate residues are conserved in the barley enzymes AXAH-I and AXAH-II (Lee et al., 2001) and are present in the tomato and rice protein sequences.
AtASD1 and AtASD2 are differentially expressed during development
To investigate the expression profiles of AtASD1 and AtASD2, RT-PCR, northern blot analysis and promoter-GUS reporter gene expression in transgenic plants were used. RT-PCR analysis was used to obtain a broad overview of gene expression. RNA from whole seedlings or tissues such as roots, rosettes, stems, flowers, and siliques from plants of different ages, was analysed for AtASD1 and AtASD2 expression. Using RT-PCR, the AtASD1 transcript was detected at high levels in all tissues examined, only decreasing slightly in older rosettes (Fig. 4). By contrast, negligible AtASD2 transcript was detected in 7, 14 and 21-d-old roots or leaves. In aerial tissues, however, AtASD2 transcript levels were higher in flowers, siliques and to a lesser extent in stem tissue (Fig. 4).
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To investigate the temporal and tissue-specificity of gene expression further, transgenic lines carrying AtASD1 and AtASD2 promoter::GUS reporter constructs were generated. Two upstream fragments of AtASD1 and AtASD2, 2.8 kb and 2.0 kb in size, respectively (Fig. 1), were fused to the GUS gene in the binary vector pBI101. These fragments each contained the intron upstream of the predicted start codon. The constructs, denoted AtASD1::GUS and AtASD2::GUS, were transformed into wild-type (Col-0) plants. From primary transformants, more than 20 lines homozygous for a single transgene locus were identified and tested for GUS expression. Although the intensity of GUS activity varied across the lines isolated, temporal and spatial expression patterns were consistent (data not shown). Two plant lines for each construct were chosen for further analysis.
In seedlings AtASD1::GUS activity was present in all tissues (Fig. 5), with greater intensity shown in the zone for cell division and expansion in the primary root apex (Fig. 5A) and emerging lateral roots (Fig. 5B), in newly differentiating leaves (Fig. 5C) and the vascular system as a whole (Fig. 5A–D). By contrast, GUS activity in AtASD2::GUS seedlings was confined to the vasculature of older root tissue (Fig. 5H), where staining was discontinuous and non-uniform (Fig. 5I). AtASD2::GUS root expression was not evident in the primary root apex or emerging lateral roots (Fig. 5J,K). Faint staining in the vascular system of young leaves and the hypocotyl was occasionally evident (not shown),
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In adult plants, AtASD1::GUS activity was observed in vegetative tissues such as mature and senescing leaves (Fig. 5F, G), trichomes (Fig. 5E) and cauline leaves (Fig. 6D). During floral development, AtASD1::GUS activity was observed in expanding sepals of young flowers (Fig. 6A), but was most evident in mature and ageing flowers (Fig. 6B, C). AtASD1 expression was observed in sepals (Fig. 6G), petals (Fig. 6H), stamen filaments (Fig. 6I) and developing siliques (Fig. 6E, F), with staining prominent in the vasculature of these tissues. AtASD1 expression was also prominent in zones of cells associated with floral abscission, both in proximal tissues at the silique base (Fig. 6F) and sites of detachment in organs such as sepals (Fig. 6G), petals and stamen filaments (not shown). AtASD2::GUS expression in adult plants was restricted to flowers that are fully developed (Fig. 6L) and was localized to the tips of anthers (Fig. 6J), petal blades (Fig. 6L, K), the floral abscission zones at the base of siliques (Fig. 6O), and in silique replum tissue (Fig. 6M, N). In contrast to AtASD1, AtASD2::GUS expression was absent in the vascular system of floral tissues, with expression observed occasionally in the vasculature of stems (Fig. 6P). In summary, AtASD1 and AtASD2 portray distinct expression profiles during development, although interestingly, both genes are expressed in floral abscission zones.
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Plant hormones and leaf senescence do not affect AtASD1 and AtASD2 expression
Plant hormones are known to be key regulators of expression of many cell wall modification enzymes. To explore the possibility that plant hormones influence AtASD1 and AtASD2 gene expression, 7 d seedlings were incubated for 24 h with each of the following; indole acetic acid, 2,4-dichlorophenoxyacetic acid, gibberellic acid, abscisic acid, kinetin, and epibrassinolide at concentrations of 0.1 µM, 1.0 µM and 10 µM. Relevant controls were included. Northern analysis demonstrated that transcript levels for AtASD1 remained high and unvarying for all treatments, while AtASD2 transcript levels remained low and unchanging (data not shown). In a parallel study, transgenic AtASD1::GUS and AtASD2::GUS seedlings were treated as described above and analysed using GUS histochemistry. Hormone-treated seedlings exhibited levels of GUS activity indistinguishable from stained control seedlings (data not shown), which coincides with the northern data.
In an additional study, AtASD1 and AtASD2 expression during leaf morphogenesis and leaf senescence was investi gated. Ageing rosettes (38 d) were dissected and leaves assigned to six groups, representing a developmental progression from very young leaves to those undergoing senescence. Total RNA was extracted and used for northern analysis. Again, AtASD1 transcript levels remained high and invariable across all samples (data not shown). This indicates that AtASD1 expression is not regulated during leaf morphogenesis or senescence. AtASD2 transcript was not detected in any samples (data not shown), which is in accordance with AtASD2 RT-PCR data for rosettes older than 2 weeks.
Possible functional roles for AtASD1 and AtASD2
To date, little is known regarding the function of plant -L-arabinofuranosidases and their regulation. In barley, it is suggested that AXAH-I and AXAH-II mediate arabinoxylan metabolism during grain germination (Ferré et al., 2000; Lee et al., 2001) and may alter the arabinoxylan component of cell walls during general cell growth processes (Gibeaut and Carpita, 1991). In tomato, three -L-arabinofuranosidase isozymes have been isolated from pericarp tissue (Sozzi et al., 2002). These are implicated in fruit ontogeny and show differential responses to gibberellic acid, 2,4-dichlorophenoxyacetic acid and ethylene. Immunolocalization studies of RG-I-associated (1->5)-arabinan epitopes have identified different developmental patterns of epitope localization in carrot and Arabidopsis root apices. In carrot root apices, the actively dividing cells of the meristem centre contain high levels of (1->5)-arabinan. The epitope is significantly reduced in surrounding tissues, including the files of elongated cells emerging from this zone. This developmental pattern is reflected in suspension-cultured carrot cells. When induced to elongate, (1->5)-arabinan-containing cell walls of proliferating cells become significantly depleted in this epitope (Willats et al., 1999). By contrast, the cell proliferation zones in Arabidopsis root apices lack the (1->5)-arabinan epitope (Willats et al., 2001a) while (1->5)-arabinan is apparent in the elongated cell files of the root and occasionally in the root cap.
AtASD1::GUS activity is intense in root apices, which may correlate with the general absence of (1->5)-arabinan in this region. AtASD1::GUS activity, and to a lesser extent AtASD2::GUS activity, is also apparent in the vasculature of roots, where other glycoside hydrolases, such as AtXyn1 and AtBXL1, are expressed (Suzuki et al., 2002; Goujon et al., 2003). These xylose-targeting enzymes are reported to have potential roles in morphogenesis and secondary cell wall formation in xylem vessels. In situ hybridization studies would aid in confirming a correlation between developmental processes, such as xylogenesis, and AtASD1 and AtASD2 expression.
Abscission involves the co-ordinated dissolution of the cell wall matrix in a zone of specialized cells, resulting in cell separation. Cell wall hydrolytic enzymes such as ß-1,4-glucanases (del Campillo et al., 1990; Lashbrook et al., 1994, 1998; del Campillo and Bennett, 1996), polygalacturonases (PG) (Hadfield and Bennett, 1998; Hong et al., 2000; Torki et al., 2000; Patterson, 2001; Sander et al., 2001; González-Carranza et al., 2002) and also expansins (Cho and Cosgrove, 2000), have been implicated in abscission processes. In this study, expression of both AtASD1 and AtASD2 in floral abscission zones is apparent. During abscission, the pectin-rich middle lamella that acts to cement adjoining cell walls is solubilized (Sexton, 1982; Taylor et al., 1993; Roberts et al., 2000). As structural L-arabinose-containing glycans are primarily associated with the pectic RG-I and RG-II polymers, it is conceivable that these glycans are metabolized during cell separation processes. This is supported by biochemical evidence that pectin-associated arabinan linkages with cellulose-hemicellulose complexes may be important for intercellular attachment (Fu and Mort, 1997; Femenia et al., 1999; Iwai et al., 2001).
Concluding remarks
In this study, two genes (AtASD1 and AtASD2) encoding putative -L-arabinofuranosidases have been described in Arabidopsis thaliana. Gene identities were assigned on the basis of sequence similarity to plant and microbial sequences classified as family 51 glycoside hydrolases. RT-PCR and GUS reporter gene analysis clearly illustrate AtASD1 and AtASD2 have different expression patterns and may have multiple roles during development. High levels of AtASD1 expression coincides with many developmental processes such as cell proliferation and vascular development during vegetative growth and the morphogenesis, senescence and abscission of floral organs. AtASD2 expression is less evident during development. In seedlings, AtASD2 expression is almost exclusively limited to the vascular system of older primary root tissue. In adult plants, AtASD2 expression is evident in some organs of mature flowers, their associated abscission zones, silique replum, and the vasculature of stems. Whilst this study has provided insights into the expression profiles of AtASD1 and AtASD2 in Arabidopsis, further investigation into substrate specificities and mutant characterization would assist in elucidating their specific biological functions.
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Acknowledgement |
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We thank Quentin Lang for excellent technical assistance with photography.
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References |
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