From the Department of Plant Science, University of Adelaide, Waite Campus, Glen Osmond, South Australia 5064, Australia
Received for publication, October 17, 2002, and in revised form, December 1, 2002
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ABSTRACT |
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An
Heteroxylans are major constituents of cell walls in the Poaceae,
which include many commercially important cereals and pasture grasses.
In the endosperm of barley grains and in elongating coleoptiles, these
polysaccharides may comprise 20-70% by weight of the walls (1) and
consist of a backbone of (1 As observed with many wall components in higher plants, the
arabinoxylans of primary cell walls can be restructured during normal
growth and development. For example, newly synthesized arabinoxylans in
maize coleoptiles are deposited in the walls in a highly substituted
form, but arabinofuranosyl residues are removed later, and this leads
to significant changes in the physicochemical properties of the
polysaccharides and hence in the walls themselves (5, 6). Removal of
Most Here we have purified a bifunctional family 3 Materials--
DEAE-cellulose (DE52) was from Whatman.
CM-Sepharose CL-6B, Polybuffer Exchanger 94, Polybuffer 96, Polybuffer
74, and molecular mass marker proteins were from Amersham Biosciences.
Bio-Gel P-100 was from Bio-Rad Laboratories. Phenylmethylsulfonyl
fluoride, bovine serum albumin, 4NPA,
4'-nitrophenyl- Enzyme Purification--
Barley (Hordeum vulgare L. cv. Clipper) (3 kg dry weight) was surface sterilized in 0.2% (w/v)
AgNO3, washed successively with sterile water, 0.5 M NaCl, and sterile water, and steeped for 24 h in
sterile water containing 100 mg/ml chloramphenicol, 100 mg/ml neomycin,
100 units/ml penicillin G, and 100 units/ml nystatin. Germinated grains
were maintained at 40-45% (w/w) moisture content by regular
application of fresh antibiotic solution for 5 days in the dark at
22 °C. Bacterial or fungal contamination of the grains was not
evident at any stage during this period. The germinated barley material
was stored overnight at Enzyme Assays--
Activities of
Protein Determination and PAGE--
Protein contents of pooled
fractions and purified proteins were measured using the Coomassie
Brilliant Blue reagent. Purity of column fractions and purified
proteins was assessed by SDS-PAGE (22) on 12.5% (w/v) polyacrylamide
gels and stained with Coomassie Brilliant Blue R-250.
Substrate Specificities--
Aryl glycosides were
used as substrates with purified proteins in standard assays at a final
concentration of 2.5 mM. Rates of hydrolysis of polymeric
substrates, at a final concentration of 0.2% (w/v), were determined
from the increase in reducing sugars (23, 24). Oligosaccharide and
monosaccharide products liberated during enzymic hydrolyzes were
analyzed by thin layer chromatography on silica gel plates (Merck).
Plates were developed in ethyl acetate:acetic acid:water (3:2:1 v/v),
and sugars were detected using the orcinol reagent (21).
Kinetic Analyses--
Kinetic parameters of purified enzymes
were determined against 4NPA and 4NPX in a concentration range of
0.25-7.0 mM. Assays were performed in triplicate, in 50 mM sodium acetate buffer, pH 4.7, containing 4 mM sodium azide and 160 µg ml Amino Acid Sequence Analysis--
NH2-terminal
sequence analyses of proteins and peptides generated by CNBr, trypsin,
or Lys-C and purified by reversed phase HPLC were performed on a
Hewlett-Packard G1005A automated protein sequencer, using Edman
degradation chemistry. Phenylthiohydantoin-derivatives were identified
by reversed phase HPLC using a Hewlett-Packard HPLC 1090 system.
To identify the COOH terminus of ARA-I, the purified enzyme was
subjected to extensive amino acid sequence analysis, as follows. First,
ARA-I was reduced with 5 mM dithiothreitol, 6 M
guanidine HCl in 50 mM Tris-HCl buffer, pH 8.5, at 65 °C
for 30 min and alkylated with 20 mM iodoacetamide for 20 min at 20 °C. Dithiothreitol was added to the alkylation solution to
remove excess iodoacetamide. Second, the reduced, alkylated protein was
fragmented with trypsin or Lys-C. Prior to trypsin digestion, buffer
exchange into 100 mM Tris-HCl buffer, pH 7.0, containing 4 M urea and 2 mM dithiothreitol, was performed
using a Nanosep Centricon (PALL Life Sciences, Ann Arbor, MI) with a
3-kDa cutoff. After diluting the ARA-1 four times with water, trypsin
(Promega) was added for 16 h at 20 °C. For Lys-C digestion, the
buffer exchange was into 25 mM Tris-HCl buffer, pH 7.0, containing 4 M urea. Digestion with endoproteinase Lys-C
(Promega) was performed for 16 h at 20 °C. The reactions were
stopped by adjusting the pH to 3 with 2% (v/v) trifluoroacetic acid.
The Lys-C and tryptic peptides were separated on a Vydac C18 protein
column (250 × 2.1 mm, 5 µm; Hesperia, CA). The eluents were (A)
(0.05% (v/v) trifluoroacetic acid and (B) 0.04% (v/v) trifluoroacetic
acid in 70% (v/v) acetonitrile. The flow rate was 0.2 ml/min, and the
gradient was composed of 2-70% (v/v) B for 90 min and 70-100% (v/v)
B for an additional 10 min. The column temperature was 40 °C, and
protein was detected by absorbance at 214 and 280 nm. Where necessary
to separate poorly resolved peptides, selected fractions were
rechromatographed using a shallower gradient. Some peptides were
fragmented further with 100 mM CNBr in 0.1 M
HCl for 16 h at 20 °C.
Mass Spectrometry--
The ARA-I enzyme was desalted by reversed
phase HPLC on a 200 × 2.1 mm POROS 20R1 column (Applied
Biosystems) with 1% (v/v) formic acid and a 5-100% (v/v)
acetonitrile gradient at 0.5 ml min
Where no mass was found for a peptide by MALDI-TOF, mass determinations
were carried out with an ABI Sciex API 300 electrospray ionization
quadrupole mass spectrometer equipped with an ion spray ion source (PE
Sciex, Thornhill, Ontario, Canada). Positive ion mass spectra were
recorded in a range from m/z 200 to 2,600 and were processed to determine the most probable molecular masses of the
peptides using Bio-Multiview software version 1.3 (PE Sciex).
RNA Isolation and PCR--
Total RNA was isolated from root,
leaf, coleoptile, and scutellum tissue from barley seedlings, 4 days
after germination, using the TRIZOL reagent (Invitrogen) as
recommended by the manufacturer. First strand cDNA was prepared
from 3 µg of total RNA using the 3'-RACE primer
(5'-GACTCGAGTCGACATCGAT17-3') (26) and the
THERMOSCRIPT reverse transcriptase system (Invitrogen).
cDNA fragments corresponding to sequences predicted from the
purified proteins were amplified from the root, shoot, and scutellum
single-stranded cDNA preparation by PCR in a mixture containing
Taq DNA polymerase, standard PCR buffer, 5 mM
dNTPs, 10% (v/v) dimethyl sulfoxide, and 1.5 mM
MgCl2, and primed with degenerate oligonucleotide primer
pairs, designed on the basis of tryptic peptide sequences. The
sequences of these primers were 5'-GGNATHCCNGCNTAYGARTGGTGG-3'
(upstream primer) and 5'-AAIGGIGGYTGRAAIGTRTCRTC-3' (downstream primer;
I is deoxyriboinosine) corresponding with ARA-I tryptic peptides, and
5'-GTNCCNGCNTAYAAGTGGTGG-3' (upstream primer) and
5'-TCYTGRTTYTGRTCIARICCCAT-3' (downstream primer) corresponding to XYL
tryptic peptides. The PCR cycles (35) consisted of a denaturation step
(94 °C, 40 s), annealing (50 °C, 40 s), and extension
(72 °C, 2 min). Products from the PCR were purified from agarose
gels following electrophoresis, and after sequence confirmation, were
used to probe cDNA Isolation--
A Nucleotide Sequence Analysis--
Both strands of isolated
cDNAs were sequenced using the dideoxynucleotide chain termination
procedure (28). Data from automated sequencing were compiled and
analyzed using the Seq-Ed program (Applied Biosystems), and further
analyses of DNA sequences and data base searches were performed using
the University of Wisconsin Genetics Computer Group software (29) in
the ANGIS suite of programs at the University of Sydney
(www.angis.org.au/WebANGIS/). Sequences were aligned using the
ClustalW program (www2.ebi.ac.uk/clustalw).
Reverse Transcription-PCR--
cDNA samples were prepared
from 3 µg of total RNA from developing grains (6 days postanthesis),
leaf (5 days after germination), coleoptile (4 days), rootlets (5 days), aleurone (3 days), and scutella (3 days). Reverse transcriptase
reactions were primed with (dT)17 primer. PCRs contained 1 µl of cDNA and 0.1 µg of each gene-specific primer. DNA
fragments corresponding to each gene were amplified by 30 cycles of PCR
consisting of 94 °C, 40 s; 55 °C, 40 s; 72 °C,
30 s. Amplified products were detected by gel electrophoresis, and
DNA bands were observed under ultraviolet light. For each set of
primers, amplified DNA was excised from the agarose gel and purified
for subsequent DNA sequence analysis. DNA sequences, in every case,
exactly matched the DNA sequence of the respective cDNA clone.
Genetic Mapping of ARA-I and XYL Genes--
Barley mapping
populations and parental lines were screened at high stringency with
probes corresponding to each of the XYL and ARA-I cDNAs, as
described (30). The DNA probe for ARA-I was the 2,400-bp 3'-end RACE
fragment, and for XYL was the 1,800-bp 3'-end RACE fragment. Filters
were hybridized with radiolabeled DNA probes at 65 °C and washed in
0.1 × SSC, 0.1% (w/v) SDS at 65 °C to remove nonspecifically
bound probe DNA. Chromosomal locations for the ARA-I and
XYL genes were allocated by correlation with genetic markers
using the Mapmaker and JoinMap software (31, 32).
Hydrophobic Cluster Analysis and Molecular
Modeling--
Hydrophobic cluster analysis (HCA) was performed using
DrawHCA (33; www.lmcp.jussieu.fr/~callebau/hca_method.html). The
three-dimensional molecular models of ARA-I and XYL were built using
the three-dimensional structure of barley Purification of Barley ARA-I and XYL--
Preliminary experiments
showed that
Two peaks of
In view of the apparent activity of the ARA-I on both 4NPA and 4NPX
during the purification process, particular care was taken to evaluate
the purities of the final enzyme preparations. The final ARA-I
preparation appeared as a single protein of molecular mass 67 kDa after
SDS-gel electrophoresis (Fig.
2A) and had an isoelectric
point of 5.5. Although the NH2-terminal sequence could not
be obtained with ARA-I, presumably because the NH2-terminal residue was blocked to Edman degradation, the sequences of 10 tryptic
peptides from the enzyme were determined. For the final XYL
preparation, which had a molecular mass of 67 kDa (Fig. 2B) and an isoelectric point of 6.7, a minor protein band greater than 100 kDa could be seen but was not resolved by additional chromatography
steps (data not shown). An NH2-terminal sequence of 38 amino acid residues was obtained for XYL, together with the sequences
of six tryptic peptides. For both ARA-I and XYL, amino acid sequence
comparisons with protein sequences in the data bases revealed that the
enzymes were members of the family 3 group of glycoside hydrolases
(13).
Kinetic Analyses--
The kinetic parameters for ARA-I and XYL
using 4NPA and 4NPX, respectively, are shown in Table
II. The Km values
for ARA-I against aryl glycosides revealed that this enzyme had
a relatively low Km for 4NPX compared with 4NPA,
although the catalytic rate was higher on 4NPA (Table II).
Comparison of kinetic parameters for XYL against 4NPA and 4NPX
demonstrated a definite preference for 4NPX (Table II). XYL had a
relatively low Km value for 4NPX (1.7 ± 0.04 mM) compared with the Km value
for 4NPA (24.8 ± 0.04 mM), the catalytic rate constant for 4NPX was twice that for 4NPA, and the catalytic efficiency factor for 4NPX was about 30 times that measured for 4NPA (Table II).
Substrate Specificities--
The pH optimum for both ARA-I and XYL
was 4.7 (data not shown). The preferred aryl glycoside substrates for
ARA-I and XYL were 4NPA and 4NPX, respectively. ARA-I could also
hydrolyze 4NPX, 4NP-
Of the polysaccharides examined, ARA-I and XYL hydrolyzed only
arabinoxylan, but hydrolytic rates were very low, and activity could be
detected only after prolonged incubation of the enzymes with this
substrate (Fig. 3A). When
ARA-I was incubated with the arabinoxylan, small amounts of both
L-arabinose and D-xylose were released, but XYL
released only D-xylose (Fig. 3A). Neither enzyme hydrolyzed larch arabinogalactan,
(1
In the presence of (1 Isolation of cDNAs--
Degenerate oligonucleotide primers
were designed to correspond with ARA-I tryptic amino acid sequences,
and PCR was performed using all combinations of forward and reverse
primers with reverse transcribed, 3-day-old barley seedling mRNA.
The nucleotide sequence of a 500-bp PCR product corresponded exactly
with the amino acid sequences of various tryptic peptides from purified
ARA-I, and the PCR product was therefore used to probe a cDNA
library from 24-48 h gibberellic acid3-induced aleurone
layers. Of 200,000 cDNA clones screened, a single 1,900-bp ARA-I
cDNA was identified. The missing 5'-end fragment of the cDNA
was subsequently isolated during additional screening of the cDNA
library, and 3'-RACE was used to generate the missing 3'-end fragment.
The fragments were used to assemble the full nucleotide sequence of the
ARA-I cDNA, and a similar strategy was used to assemble a near
full-length cDNA encoding the XYL enzyme (data not shown). In both
cases a strong bias in codon usage was evident for the two genes, with G or C residues found in the wobble base position of about 95% of
ARA-I and XYL codons in the region encoding the mature enzymes (data
not shown).
Primary Structures of the Enzymes--
The complete amino acid
sequences of the barley ARA-I and XYL enzymes were deduced from the
nucleotide sequences of the corresponding cDNAs (Fig.
4). For the XYL enzyme, 38 amino acids
were sequenced from the NH2 terminus of the purified
enzyme. This showed that the NH2-terminal residue of the
mature enzyme was the Ala residue of the ADPPF sequence indicated in
Fig. 4. The sequence of the 38 NH2-terminal amino acid
residues corresponded with the sequence deduced from the cDNA in 31 positions, whereas the sequences of another 168 amino acids derived
from 8 tryptic peptides matched exactly with the sequence deduced from
the cDNA. The fact that the experimentally determined and deduced
NH2-terminal sequences did not exactly match raised the
possibility that the cDNA sequence was in fact a composite sequence
of genes encoding two closely related XYL isoenzymes from the barley
cDNA libraries. However, the overlapping region of the two 5'
cDNA fragments was 861 bp in length (Fig. 4), and the overlapping
nucleotide sequences of the various cDNA fragments matched exactly.
At this stage we are unable to explain this apparent discrepancy.
No NH2-terminal sequence was obtained for the purified
ARA-I enzyme, although the sequences of a total of 126 amino acid
residues from 9 different tryptic peptides matched the sequence deduced from the cDNA fragments (Fig. 4). A few amino acid differences were
observed, but these were attributed to differences in the varieties
used to isolate the enzyme and the cDNA library. The NH2-terminal sequence of ARA-I (AEAQAQAPVF) was predicted
using the SigCleave program and corresponded to the experimentally
determined sequence for the NH2 terminus of the XYL enzyme
(Fig. 4). In both cases the
If it is accepted that the NH2-terminal residue of ARA-I is
as indicated in Fig. 4, the ARA-I and XYL cDNAs both encode mature polypeptides of 748 amino acids, and these show 51% positional identity for the two enzymes. Signal peptides of 29 residues (Fig. 4)
were detected for each enzyme, and these have characteristics typical
of those from other eukaryotic signal peptides that direct nascent
polypeptides to the endoplasmic reticulum (44). The molecular mass
calculated from the deduced amino acid sequence was 79,184 Da for
ARA-I, which had a calculated isoelectric point of 5.7. The
corresponding values calculated for XYL were 80,500 Da and 6.5 for XYL,
respectively. Although the calculated isoelectric point values
correspond well with the values of 5.5 and 6.7 for ARA-I and XYL,
respectively, as determined from the purified enzymes, the molecular
masses deduced from the cDNA sequences are considerably higher than
the values of 67 kDa obtained for the purified enzymes on SDS gels run
under reducing conditions (Fig. 2). This would suggest that processing
of the primary translation product might have occurred and that a
peptide fragment of up to about 12 kDa might have been removed during
enzyme maturation. However, the NH2-terminal end of XYL has
certainly not been processed, and the presence of a Lys-C peptide
starting at residue 29 of ARA-I suggests that a fragment of 12 kDa has
not been cleaved from the NH2 terminus of this enzyme
either (Fig. 4). Internal processing of the enzymes would also be
possible, but the 5 mM 2-mercaptoethanol included here in
the gel loading buffer during electrophoresis and during enzyme
purification would dissociate individual peptide chains linked by
disulfide bonds (45), and we could find no evidence for internal
processing of the two enzymes.
In view of the discrepancy between observed and predicted apparent
molecular mass values, ARA-I and XYL were examined by MALDI-TOF mass
spectrometry. Broad peaks of 69.4 kDa and 68.2 kDa, with widths of 3-5
kDa, were obtained for ARA-I and XYL, respectively (data not shown).
The amino acid sequence of ARA-I was investigated in more detail. Using
a combination of MALDI-TOF and electrospray ionization mass
spectrometry of tryptic and Lys-C peptides, coupled with Edman sequence
analysis of the peptides, most of the NH2-terminal and
central peptides predicted from the ARA-I cDNA sequence could be
identified (Fig. 4). However, the COOH-terminal region of the enzyme
predicted by the cDNA sequence was not detected in any of the
enzymic digests, and it was concluded on this basis and on the basis of
domain prediction (46) that the actual COOH terminus of the mature
ARA-I enzyme is in the vicinity of Met-614 (Fig. 4).
Catalytic Amino Acid Residues and Active Sites--
Based on
multiple sequence alignments (47) and HCA (33), the putative catalytic
nucleophiles are predicted to be Asp-275 for ARA-I and Asp-268 for XYL
(Fig. 4). These residues are absolutely conserved in family 3 glycoside
hydrolases (19). Prediction of the catalytic acid/base residues of the
enzymes is somewhat more complicated. Multiple sequence alignments and
HCA clearly identify two candidate amino acid residues for this role
(data not shown). Hrmova et al. (48) suggested that the
catalytic acid/base for
Other features of family 3 glycoside hydrolases which can
be observed in the ARA-I and XYL sequences include the conserved WGR
and KH motifs, beginning at residues Trp-147 and Lys-192 for ARA-I, and
Trp-139 and Lys-185 for XYL. These motifs are probably involved in
substrate binding (34, 48). In addition, sequences similar to the
conserved COOH-terminal antiparallel loop of family 3 enzymes (34) were
present in the region starting at about residue 559 in ARA-I and 562 in
XYL (Fig. 4).
Expression Analysis of ARA-I and XYL Genes--
Transcript levels
of ARA
ARA-I transcripts were detected in developing grains and in the
vegetative tissues of rootlets, coleoptiles, and leaves (Fig. 5). ARA-I does not appear to be
transcribed in aleurone or scutellum tissue 3 days after germination.
XYL transcripts were found in each of the barley tissues and at
relatively high levels (Fig. 5).
Genetic Mapping of ARA-I and XYL Genes--
Single dominant
bands were evident when Southern hybridization analyses of parental
lines and mapping population DNA samples were probed with ARA-I and XYL
cDNAs (data not shown). This suggested that single genes encoding
these enzymes are present on the barley genome, although it should be
noted that the hybridizations were performed at high stringency to
avoid cross-hybridization between the ARA-I and XYL probes, which are
about 50% identical, and related genes might therefore have gone
undetected. Indeed, preliminary amino acid sequence analysis of a
protein band enriched in ARA-II revealed differences with ARA-I
sequences. Despite the fact that ARA-II was not completely purified, it
could be concluded from the sequence comparisons that there are at
least two genes encoding
Restriction fragment length polymorphisms (RFLPs) for the DNA probes
were rare, with only one RFLP for ARA-I found in DNA digested with
HindIII from the parents Chebec and Harrington and for XYL
RFLPs were present only for Clipper and Sahara genomic DNA digested
with EcoRI or DraI. The ARA-I gene is located on the long arm of barley chromosome 2H, between the molecular markers ABC165 and BCD512, and XYL is found near the centromere of barley chromosome 6H, between markers Bmag9 and BCD269 (Fig.
6).
An Using amino acid sequences generated from the purified barley ARA-I and
XYL enzymes, several cDNAs were isolated, and near full-length
cDNA sequences were subsequently assembled (Fig. 4). Deduced amino
acid sequences indicated that both enzymes have a typical endoplasmic
reticulum-targeting signal peptide (Fig. 4) that presumably directs
secretion from cells in which they are synthesized. This is a
significant observation, given recent indications that the
(1 Although the NH2 terminus of ARA-I could not be defined
with certainty, the cDNAs encode primary translation products of
748 amino acid residues (Fig. 4). The calculated molecular masses of
the enzymes, based on these deduced amino acid sequences, are about 80 kDa. This value is much higher than the apparent molecular mass values
of 67 kDa observed during SDS-gel electrophoresis of the purified
enzymes (Fig. 2) and represents a much longer polypeptide than other
plant members of the family 3 group of glycoside hydrolases (19). Mass
spectrometry was therefore used to examine further the molecular masses
of the two enzymes and in both cases confirmed that the enzymes were
~67 kDa in size. Particular attention was paid to ARA-I, for which
the analysis of proteolytic peptides accounted for all regions of the
enzyme except the COOH terminus predicted from the cDNA sequence
(Fig. 4). The amino acid sequence data also suggested that the COOH termini of the enzymes were heterogeneous and that a single
COOH-terminal residue could therefore not be identified. At this stage
the weight of evidence suggests that the COOH termini of both ARA-I and
XYL are close to the Met-614/Tyr-606 residue of ARA-I/XYL, respectively (Fig. 4). Thus, more than 130 amino acid residues appear to have been
removed from the COOH termini during post-translational processing of
the enzymes. These values may be compared with the 605 residues found
in the family 3 barley Comparison of the amino acid sequences of the mature enzymes with other
members of the family 3 glycoside hydrolases suggested that
COOH-terminal processing does not occur in all members of the family.
Although The relatively relaxed substrate specificities observed here for the
family 3 ARA-I and XYL enzymes from barley can be rationalized in terms
of their predicted three-dimensional structures. The three-dimensional
structure of a family 3 The barley -L-arabinofuranosidase and a
-D-xylosidase, designated ARA-I and XYL, respectively,
have been purified about 1,000-fold from extracts of 5-day-old barley
(Hordeum vulgare L.) seedlings using ammonium sulfate
fractional precipitation, ion exchange chromatography,
chromatofocusing, and size-exclusion chromatography. The ARA-I has an
apparent molecular mass of 67 kDa and an isoelectric point of 5.5, and
its catalytic efficiency during hydrolysis of 4'-nitrophenyl
-L-arabinofuranoside is only slightly higher than during
hydrolysis of 4'-nitrophenyl
-D-xyloside. Thus, the
enzyme is actually a bifunctional
-L-arabinofuranosidase/
-D-xylosidase. In
contrast, the XYL enzyme, which also has an apparent molecular mass of
67 kDa and an isoelectric point of 6.7, preferentially hydrolyzes
4'-nitrophenyl
-D-xyloside, with a catalytic efficiency ~30-fold higher than with 4'-nitrophenyl
-L-arabinofuranoside. The enzymes hydrolyze wheat flour
arabinoxylan slowly but rapidly hydrolyze oligosaccharide products
released from this polysaccharide by
(1
4)-
-D-xylan endohydrolase. Both enzymes
hydrolyze (1
4)-
-D-xylopentaose, and ARA-I can
also degrade (1
5)-
-L-arabinofuranohexaose. ARA-I
and XYL cDNAs encode mature proteins of 748 amino acid residues which have calculated molecular masses of 79.2 and 80.5 kDa,
respectively. Both are family 3 glycoside hydrolases. The discrepancies
between the apparent molecular masses obtained for the purified enzymes and those predicted from the cDNAs are attributable to
COOH-terminal processing, through which about 130 amino acid residues
are removed from the primary translation product. The genes encoding
the ARA-I and XYL have been mapped to chromosomes 2H and 6H,
respectively. ARA-I transcripts are most abundant in young roots, young
leaves, and developing grain, whereas XYL mRNA is detected in most
barley tissues.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
4)-
-linked
D-xylopyranosyl residues substituted predominantly with
-L-arabinofuranosyl residues. The
-L-arabinofuranosyl residues can be linked to O-3, O-2,
or both O-3 and O-2 of xylanopyranosyl residues of the
(1
4)-
-D-xylan backbone, and other substituents or
short side chains are also detected in low abundance (2, 3). The
-L-arabinofuranosyl residues can be esterified with
hydroxycinnamic acids, in particular ferulic acid, which may
form cross-bridges between adjacent arabinoxylan chains, or with
lignin, by oxidative dimerization (4).
-L-arabinofuranosyl residues is also observed when wall
arabinoxylans are degraded (7, 8). The presence of
-L-arabinofuranosidases in germinated barley grain or in
isolated aleurone layers has been taken as evidence that these enzymes
perform this function during the mobilization of the starchy endosperm
after cereal grain germination (9, 10), although this activity has also
been attributed to a separate group of enzymes, known as arabinoxylan
-L-arabinofuranohydrolases (11, 12).
-L-arabinofuranosidases are so designated because
they can hydrolyze the synthetic aryl glycoside 4'-nitrophenyl
-L-arabinofuranoside (4NPA),1 and although they
are presumed to be responsible for changes in arabinoxylans during wall
modification or degradation, this class of enzymes may be subdivided
into several quite distinct groups. Thus,
-L-arabinofuranosidases have been classified in glycoside hydrolase families 3, 43, 51, 54, and 62 (13;
afmb.cnrs-mrs.fr), and members of each family exhibit characteristic
substrate specificities, action patterns, and reaction mechanisms
(14-19), and three-dimensional structures (20). Although most of the
characterized enzymes are from saprophytic or rumen microorganisms,
several plant
-L-arabinofuranosidases have also been
identified. Family 51 arabinoxylan arabinofuranohydrolases, which
remove
-L-arabinofuranosyl residues from polymeric
arabinoxylans, have been purified from germinated barley grain, and
their primary structures have been defined (11, 12). There are other
reports of the purification or partial purification of higher plant
-L-arabinofuranosidases, but in most cases no amino acid
sequence information is available, and it is therefore not possible to
classify the enzymes accurately, to draw conclusions about their
reaction mechanisms, or to identify their true substrates and
biological functions.
-L-arabinofuranosidase/
-D-xylosidase
(ARA-I) from young barley seedlings, defined its kinetic and enzymic
properties, and determined its complete amino acid sequence from
corresponding cDNAs. The enzyme is unable to hydrolyze
arabinoxylans at a significant rate but could play an important role in
the complete depolymerization of arabinoxylans through its ability to
hydrolyze oligosaccharides released from the polysaccharide by
(1
4)-
-D-xylan endohydrolases. In parallel, a
family 3
-D-xylosidase (XYL) was purified and characterized.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-xylopyranoside (4NPX), all other aryl
glycosides, laminarin, and arabinogalactan were from Sigma. Coomassie
Brilliant Blue reagent was from Pierce, and wheat arabinoxylan,
(1
5)-
-L-arabinofuranohexaose,
(1
4)-
-D-xylopyranopentaose, and barley
(1
3,1
4)-
-D-glucan were from Megazyme
(Bray, Ireland). Larchwood (1
4)-
-D-xylan and the
Thermomyces endoxylanase were kindly provided by Dr. Peter Biely (Institute of Chemistry, Slovak Academy of Sciences, Bratislava, Slovak Republic).
20 °C prior to homogenization at 4 °C
in 1.5 volumes of 0.1 M sodium acetate buffer, pH 5.0, containing 10 mM EDTA, 10 mM sodium azide, 3 mM 2-mercaptoethanol, and 3 mM
phenylmethylsulfonyl fluoride. Ammonium sulfate fractional
precipitation was performed as described previously (21), and the
enzyme purification procedures were as shown in Scheme
1.
View larger version (18K):
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Scheme 1.
Summary of procedures for the purification
of ARA-I and XYL.
-L-arabinofuranosidase and
-D-xylosidase
were determined spectrophotometrically using 4NPA and 4NPX,
respectively. Assays were performed at 37 °C in 50 mM
sodium acetate buffer, pH 5.0, containing 4 mM sodium azide
and 0.04% (w/v) substrate. Reactions were terminated by the addition
of 2 volumes of saturated sodium tetraborate solution. One unit of
activity is defined as the amount of enzyme releasing 1 µmol of
4-nitrophenol/min, as measured by absorbance at 410 nm.
1 bovine serum
albumin. S.E. values for assays were less than 5%. Kinetic data were
processed by a proportional weighted fit using a nonlinear regression
analysis program based on Michaelis-Menten enzyme kinetics (25).
1 at 40 °C over 15 min. For MALDI-TOF mass spectrometry, desalted ARA-1 and XYL (0.5 µl)
mixed with 1 µl of matrix solution (0.1% w/v protein in 30% v/v
acetonitrile, 1% v/v formic acid) were spotted onto a target plate and
analyzed in a Voyager-DE STR mass spectrometer (Applied Biosystems).
Peptide fractions (0.75 µl) were mixed with
-cyano-4-hydroxycinnamic acid solution (0.75 µl, 5 g/liter in 50%
v/v acetonitrile) and spotted onto a MALDI-TOF mass spectrometry target
plate, air-dried, and analyzed in a Voyager-DE STR mass spectrometer.
Experimental monoisotopic masses were compared with theoretical peptide
masses obtained from the DNA sequence, using the software MS-Digest
within the ProteinProspector tool (prospector.ucsf.edu/).
ZAP and
gt11 cDNA libraries. The 3'-ends of the
ARA-I and XYL cDNAs were amplified using the 3'-end RACE PCR
procedure (26); gene-specific primers were based on cDNA sequences
of partial cDNA clones that were isolated from cDNA libraries.
The ARA-I 3'-end cDNA was amplified by two successive rounds of PCR
(98 °C, 40 s; 50 °C, 40 s; 72 °C, 3 min) with
root/shoot/scutellum cDNA, 3'-RACE adaptor primer 5'-GACTCGAGTCGACATCG-3', and gene-specific oligonucleotide primers 5'-CGGCGTACGAGTGGTGGTCCGAAG-3' (round 1) and
5'-CGCTGCACGGCGTGTCATACGT-3' (round 2). The 3'-end of the XYL cDNA
was isolated using a single round of PCR, as described for ARA-I, with
the XYL-specific primer, 5'-GGATACATCACGTCGGAC-3'. The largest
amplified products were purified from agarose gels, ligated into the
pGEM TEasy vector (Promega) using T4 DNA ligase (New England BioLabs),
and introduced into competent DH5-
cells by electroporation using
the Bio-Rad Gene-Pulser apparatus.
ZAP-cDNA library (Stratagene,
La Jolla, CA) was prepared from poly(A)+ RNA of 24-48 h
gibberellic acid3-treated barley (cv. Clipper) aleurone
layers, and a
gt11 library was prepared from 12-day-old barley (cv.
Klages) seedlings (Clontech). The libraries were
screened on nitrocellulose membranes (Micron Separations Inc.,
Westborough, MA). Plaque replicas were hybridized with
[
-32P]dCTP-labeled ARA-I cDNA (500 bp) and XYL
cDNA (1,300 bp) fragments as described (27). Positive clones were
identified by autoradiography and purified by further rounds of
screening. The cDNA inserts were excised from the Uni-ZAP XR vector
(
ZAP clone) or, in the case of
gt11 clones, subcloned into the
pBluescript SK(+) vector (Stratagene).
-D-glucan
glucohydrolase (www.rcsb.org/pdb; PDB accession code 1IEQ; 34) as a
template, and Modeler, version 6.1 (35). The models were checked
manually on O (36) and evaluated using PROCHECK (37) and PROSAII (38).
The three-dimensional structure of
-L-arabinofuranose
was built and minimized in SYBYL 6.62 (39), and molecular surfaces of
-D-xylopyranose (PDB accession code 1B3V) and
-L-arabinofuranose were computed in GRASP (40), using a
probe size of 1.4 Å and all atoms having fixed radii.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-L-arabinofuranosidase and
-D-xylosidase activities in young barley
seedlings reached a peak 4-5 days after germination. From 3 kg of
germinated barley grain, an extract of soluble proteins from 5-day-old
barley seedlings yielded 45 and 46 units of 4NPA- and 4NPX-hydrolyzing
activities, respectively (Table I). The
steps used for the purification of the ARA-I and XYL isoenzymes are
summarized in Scheme 1.
Enzyme yields and purification factors of ARA-I and XYL
-L-arabinofuranosidase activity were
resolved on the DEAE-cellulose column at ~90 mM NaCl and
140 mM NaCl (Fig. 1A) and were designated ARA-I
and ARA-II, respectively. Attempts to purify ARA-II completely were
unsuccessful, and this isoenzyme will not be described here. The ARA-I
fractions always contained
-D-xylosidase activity at
~15% of their
-L-arabinofuranosidase activity,
throughout the purification of ARA-I (Fig. 1B) and in the
final purified enzyme preparation. It appeared therefore that although
ARA-I had a preference for 4NPA, it could also hydrolyze 4NPX. The
final ARA-I preparation was purified 1,080-fold and represented about
2% of the initial activity (Table I). It should be noted that the true
purification factor of ARA-I was likely to be significantly higher than
1,080-fold because of the ARA-II present in the initial tissue
extracts. The XYL enzyme was separated from ARA-I during the initial
DEAE-cellulose chromatography step (Scheme 1) and, after resolution
from other proteins on CM-cellulose (Fig. 1C), was
ultimately purified 960-fold (Table I).
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Fig. 1.
Chromatography of the 40-60%
(NH4)2SO4-insoluble fraction of
barley seedling extracts. A, material precipitated at
40-60% saturated (NH4)2SO4 was
applied to DEAE-cellulose at pH 8.2, and bound proteins were eluted
with a 2-liter gradient of 0-350 mM NaCl. Fractions (20 ml) were assayed against 4NPA ( ) and 4NPX (
) and protein (
).
B, fractions 21-29 from DEAE-cellulose were pooled and
applied to CM-Sepharose at pH 4.2. Bound proteins were eluted with an
800-ml gradient of 0-400 mM NaCl, and fractions were
assayed as described for DEAE-cellulose. C, material not
bound to DEAE-cellulose was concentrated and applied to CM-Sepharose at
pH 5.0. Bound protein was eluted with a 1.6-liter gradient of 0-400
mM NaCl.
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Fig. 2.
SDS-PAGE of protein fractions during the
purification of ARA-I and XYL. A: lane 1,
molecular mass markers; lane 2, seedling homogenate;
lane 3, 40-60%
(NH4)2SO4 fraction; lane
4, DEAE-cellulose ARA-I peak (fractions 21-29); lane
5, CM-Sepharose pooled fractions 10-14; lane 6,
Polybuffer Exchanger 94 fractions; lane 7, purified ARA-I
after Bio-Gel P-100. B: lanes 1-3, as for
A above; lane 4, DEAE-cellulose unbound fraction;
lane 5, CM-Sepharose pooled fractions 39-46; lane
6, Polybuffer Exchanger 94 fractions; lane 7, purified
XYL after Bio-Gel P-100.
Kinetic parameters of ARA-I and XYL
-D-galactopyranoside, and
4NP-
-L-arabinopyranoside, with 20, 16, and 11% of the
specific activity observed for 4NPA, respectively (Table
III). The preference of XYL for 4NPX was
more pronounced, with specific activities for other aryl glycosides no
greater than 3% of that for 4NPX (Table III).
Relative activities of ARA-I and XYL against aryl glycosides
3,1
4)-
-D-glucan, laminarin, or CM-xylan
(data not shown).
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Fig. 3.
Substrate specificities of ARA-I and
XYL. A, thin layer chromatography of hydrolytic
products released by ARA-I and XYL (first and second
lanes, respectively) from wheat flour arabinoxylan. The
third lane shows products released from the arabinoxylan by
the Thermomyces endoxylanase. Products of the endoxylanase
mixed with ARA-I or XYL are shown in the fourth and
fifth lanes, respectively. Standards were
L-arabinofuranose, D-xylopyranose, and
oligoxylopyranosides xylobiose-xylohexaose (sixth,
seventh, and eighth lanes, respectively).
B, products released when ARA-I and XYL were incubated with
xylopyranopentaose after 0.5, 10, and 60 min. C,
products released when ARA-I and XYL were incubated with
(1 5)-
-L-linked arabinofuranohexaose for 10 and 60 min.
4)-
-D-xylan endohydrolase,
hydrolysis rates increased dramatically, and both ARA-I and XYL
released large amounts of D-xylose; ARA-I also released
some L-arabinose (Fig. 3A). In both cases, ARA-I
or XYL was added at a concentration 1/10 of that added in assays
without the addition of the (1
4)-
-D-xylan
endohydrolase. Thus, the partial endohydrolysis of the arabinoxylan
greatly enhanced the release of oligosaccharides and monosaccharides
from this substrate (Fig. 3A). XYL completely hydrolyzed
xylopyranopentaose to D-xylose within 1 h. ARA-I was less efficient in hydrolyzing xylopyranopentaose and after 1 h, had degraded the substrate to approximately the same degree as XYL
after 10 min (Fig. 3B).
(1
5)-
-L-Arabinofuranohexaose was only partially
degraded by ARA-I in 1 h, and XYL did not hydrolyze this substrate
to any significant extent (Fig. 3C).
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Fig. 4.
Alignment of ARA-I and XYL amino acid
sequences. Identical residues in both sequences are shown in
red. The pairwise alignment was prepared using ClustalX
(41). The ARA-I and XYL cDNAs encode mature polypeptides of 748 amino acid residues with signal peptides of 29 residues each
(shading). The amino acid residues are numbered from the
NH2-terminal amino acid residues (arrowheads) of
the mature enzymes. An arrowhead is also used to indicate
the likely COOH terminus of ARA-1, and asterisks indicate
potential N-glycosylation sites (marked as CHO).
Arrows indicate the putative catalytic nucleophiles (Asp-275
for ARA-I and Asp-268 for XYL) and putative catalytic acid/bases
(Glu-481 for ARA-I and Glu-474 for XYL). Blue overlines
above the sequence of ARA-1 indicate the amino acid sequences that were
confirmed by either NH2-terminal or peptide sequencing with
MALDI-TOF or electrospray ionization quadrupole mass spectrometry
analyses, after proteolytic cleavage by Lys-C, trypsin, or CNBr.
3,
1 rule of von Heijne (42) was
satisfied, but there was no obvious reason why the
NH2-terminal residue of ARA-I would be blocked to Edman
degradation (43).
-D-xylosidase-like members of
the family 3 group would correspond to Glu-479 for ARA-I and Glu-472
for XYL. However, at this stage we believe that the catalytic residues
could just as easily be Glu-481 for ARA-I and Glu-474 for XYL (Fig. 4).
Molecular modeling experiments, in which the known three-dimensional
structure of the barley
-D-glucan glucohydrolase is used
as a template (35), indicate that the Glu-481 residue is more
appropriately positioned with respect to the known catalytic acid/base
residue Glu-491 of the
-D-glucan glucohydrolase (data
not shown). In these models the catalytic acid/base and nucleophile
amino acid residues are about 6.5 Å apart. However, it must be pointed
out that the sequence identities of the template and target enzymes are
~30% and that this is considered in the "twilight zone" of reliability of the molecular modeling programs (49).
I and XYL in various barley tissues, including developing
grain, aleurone, scutellum, rootlets, coleoptiles, and leaves, both
etiolated and light-grown, were assessed by Northern hybridization
analyses, but hybridization signals were very low. Gene-specific primers were therefore synthesized to amplify
specifically short DNA fragments, using reversed transcription PCR,
from reverse transcribed total RNA from each plant tissue sample. Sense
oligonucleotide primers corresponded to cDNA sequences within the
coding region of respective cDNAs, and antisense primers were
designed for sequences within the unique 3'-untranslated regions of
respective cDNAs.
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Fig. 5.
Reverse transcription-PCR analysis for ARA-I
and XYL. DNA was amplified from total RNA preparations of
developing barley grain (DG), aleurone (Al),
scutellum (Sc), root (R), coleoptile
(C), etiolated leaf (Let), and
light-grown leaf (Llg), using ARA-I
(A) and XYL (B) gene-specific primers.
M indicates molecular markers, and C is a
control, in which no cDNA was present.
-L-arabinofuranosidases in
barley (data not shown).
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Fig. 6.
The positions of the ARA-I
and XYL genes on barley chromosomes 2H and
6H. A, ARA-I was mapped using a Chebec and
Harrington mapping population. ARA-I is indicated with an
arrow. B, XYL was mapped using a
Clipper and Sahara mapping population. Xyl is indicated with
an arrow.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-L-arabinofuranosidase and a
-D-xylosidase, both of which are family 3 glycoside
hydrolases (13), were purified ~1,000-fold (Table I) from extracts of
germinated barley grain as outlined in Scheme 1. The purified enzymes
have been designated ARA-I and XYL, respectively. A second, less
abundant,
-L-arabinofuranosidase isoenzyme, designated
ARA-II, was detected in the extracts (Fig. 1A) but was not
purified to homogeneity. The ARA-I and XYL enzymes have apparent
molecular masses of 67 kDa (Fig. 2) and isoelectric points of 5.5 and
6.7, respectively. Examination of their substrate specificities and
kinetic properties indicated that
-L-arabinofuranosidase ARA-I can also hydrolyze 4NP-
-D-xylopyranoside,
4NP-
-D-galactopyranoside, and
4NP-
-L-arabinopyranoside at significant rates, whereas
the XYL enzyme has a more restricted, or "tighter," specificity for
-D-xylosides (Table III). Thus, the XYL enzyme
hydrolyzes not only 4NPX but also, with a 30-fold lower catalytic
efficiency, 4NPA. The catalytic efficiency factor for ARA-I was of the
same order of magnitude for 4NPA and 4NPX but slightly higher for 4NPA (Table II). For ease of expression we have referred to the enzyme here
as an
-L-arabinofuranosidase, but because ARA-I can
hydrolyze both substrates efficiently, we acknowledge that it should
probably be referred to as a bifunctional
-L-arabinofuranosidase/
-D-xylopyranosidase and that both activities might be important for its biological function
in planta. Certain family 43 (50, 51), 54 (52-54), and 62 (16)
-L-arabinofuranosidases and
-D-xylopyranosidases show similar flexibility in their
substrate specificities.
4)-
-D-xylan endohydrolase involved in
arabinoxylan depolymerization in germinated barley grain is not located
in the endomembrane secretory compartment of aleurone layers but is
found instead in the cytosol and is likely to be released from aleurone
layers only after programmed cell death (56, 57). In isolated aleurone
layers,
-L-arabinofuranosidases and
-D-xylosidases are secreted and can be detected in the
surrounding medium much earlier than the (1
4)-
-xylan
endohydrolases (10). Thus, the secretion from aleurone layers of
endohydrolases,
-L-arabinofuranosidases, and
-D-xylosidases involved in arabinoxylan degradation is
clearly not coincident.
-D-glucan glucohydrolase (58). No
biological rationale for COOH-terminal processing of the barley ARA-I
and XYL enzymes can be provided at this stage. In the case of the
barley (1
4)-
-D-xylan endohydrolase, both
NH2- and COOH-terminal processing of the primary
translation product occurs (56, 57).
-D-xylosidases from other higher plants are
similar in size to those purified here from
barley,2 the
-D-xylosidases from Aspergillus niger (59),
Aspergillus oryzae (60), and Erwinia chrysanthemi
(61) are much larger (~85 kDa) than the barley enzymes and correspond
in size to those predicted from cDNA sequences.
-D-glucan glucohydrolase from
barley has been solved (62), and although it is the only family 3 crystal structure available, it has been used to model three-dimensional structures of other family 3 enzymes (19). Molecular
modeling suggests that the barley ARA-I and XYL enzymes have overall
structures similar to that of the
-D-glucan
glucohydrolase from barley, although the three-dimensional conformation
of the 130-amino acid residue COOH-terminal region of ARA-I and XYL, which is not present in the
-D-glucan glucohydrolase
group, cannot be modeled (data not shown).
-D-glucan glucohydrolase has a broad
specificity for different linkage types in unsubstituted oligomeric and
polymeric
-D-glucan substrates (48, 63), probably
because only two glucosyl residues of the substrate enter the active
site pocket and because the glucosyl residue bound at subsite +1 is
located between two tryptophan residues that allow some positional
flexibility (48). The remainder of the substrate projects away from the enzyme surface, and activity is therefore relatively independent of
substrate shape and hence of linkage type (34). The barley ARA-I and
XYL enzymes examined here also exhibit some flexibility in substrate
specificity. Both 4NPA and 4NPX can fit in their catalytic sites. To
provide a structural rationale for this observation, the
three-dimensional structure of
-D-xylopyranose was taken from the Protein Data Bank and the three-dimensional structure of
-L-arabinofuranose was built. When the two structures
were superimposed, a similar stereochemistry was observed about C-1, C-2, and C-3 in both pentoses, and their overall hydrodynamic volumes
were also similar (Fig. 7). It is
therefore not surprising that the active site of ARA-I can accommodate
both substrates.
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Fig. 7.
Comparison of three-dimensional conformations
of -D-xylopyranose and
-L-arabinofuranose. The two
stereochemically related pentoses
-D-xylopyranose and
-L-arabinofuranose (in cpk colors) are
superimposed and show the spatial coincidence of C-1, C-2, and C-3 of
the two pentoses. Molecular surfaces of
-D-xylopyranose
and
-L-arabinofuranose are colored in white
and black, respectively, and are indicative of the
hydrodynamic volumes of the molecules. The image was generated using
GRASP (40).
In addition to the aryl glycosides, both enzymes hydrolyze linear
oligosaccharides, but neither hydrolyzes substituted polysaccharides. Similarly, the fact that no arabinose is removed from
oligoarabinoxylosides released by (1 4)-
-D-xylan
endohydrolase action (Fig. 3) suggests that neither enzyme is able to
hydrolyze substituted oligomeric substrates completely. Only
unsubstituted oligoxylosides or oligoarabinoxylosides with two to three
unsubstituted xylosyl residues at their nonreducing ends would be
expected to fit into a substrate-binding pocket of the shape found in
other family 3 enzymes (19, 48), and only xylose would be released.
Family 3 glycoside hydrolases from higher plants can be grouped into
two major clades, based on amino acid sequence alignments (48). One
group contains the broad specificity -D-glucan
glucohydrolases, and the other contains
-D-xylosidases
and
-L-arabinofuranosidases. As expected, the ARA-I and
XYL enzymes characterized here fall into the second group (data not
shown). Although the catalytic amino acid residues, corresponding to
Glu-481 and Asp-275 for ARA-I and Glu-474 and Asp-268 for XYL (Fig. 4),
are conserved in higher plant family 3 glycoside hydrolases, Hrmova
et al. (48) provided a structural explanation for the
differences in substrate specificity of the two groups. Thus, the
conserved amino acid residue Asp-95 in the
-D-glucan
glucohydrolase group that binds the C6-OH of the glycosyl residue bound
at subsite
1 is not found in the
-L-arabinofuranosidase/
-D-xylosidase
group. Clearly, the pentoses L-arabinofuranose and
D-xylose have no C6-OH group, and the
-L-arabinofuranosidase/
-D-xylosidases
have a Glu residue in the position corresponding to Asp-95 of the
-D-glucan glucohydrolase group.
When the phylogeny of the
-L-arabinofuranosidase/
-D-xylosidase
group of family 3 enzymes is examined in more detail (Fig. 8), the higher plant representatives are
clearly separated from the fungal representatives. There is one
bacterial sequence of Thermotoga neapolitana in this group
(Fig. 8). In most cases the true substrate specificities of enzymes
encoded by the genes shown in Fig. 8 have not been investigated, and
the
-D-xylosidase assignment of identity is based on
similarities with a small number of partially characterized enzymes.
The dual
-L-arabinofuranosidase/
-D-xylosidase specificity of the barley ARA-I has not been reported for other members
of family 3 (13). It is noteworthy that the barley ARA-I is some
distance from XYL in the phylogenetic tree, and this may eventually
provide clues for more detailed classification of closely related
enzymes in this family.
|
To provide some insight into the likely biological functions of the
barley ARA-I and XYL enzymes, expression patterns of the genes were
investigated, together with the action of the enzymes on well defined
oligomeric and polymeric substrates. Reverse transcription-PCR showed
the presence of XYL mRNA in all tissues examined. However, ARA-I
mRNA appeared to be absent, or in very low abundance, in the
aleurone layer and scutellum of germinated grain (Fig. 5). This is
somewhat surprising, given that -L-arabinofuranosidase activity, measured by activity on 4NPA, has been widely reported in the
media surrounding isolated barley aleurone layers (9, 10, 65). At the
substrate specificity level, XYL was able to hydrolyze
(1
4)-
-D-xylopentaose to xylose relatively quickly but exhibited no activity against
(1
5)
-L-arabinofuranohexaose (Fig. 3,
B and C). In contrast, ARA-I hydrolyzed
(1
5)
-L-arabinofuranohexaose to arabinose and
(1
4)-
-D-xylopyranopentaose to xylose, albeit at
slow rates (Fig. 3, B and C).
Neither enzyme hydrolyzed arabinoxylan at a significant rate, but both
ARA-I and XYL rapidly released xylose from oligoarabinoxylosides or
oligoxylosides that were first released from the arabinoxylan by the
action of (1 4)-
-D-xylan endohydrolase. The low
levels of arabinose in these hydrolysates (Fig. 3A) were
unexpected, given that this polysaccharide contains about 30%
(mol/mol)
-L-arabinofuranosyl residues (55), but
suggest that neither enzyme can bypass substituted xylosyl residues in
oligoarabinoxylosides. In summary, it might be concluded that the ARA-I
and XYL enzymes could participate in further hydrolysis of
oligosaccharides released from arabinoxylans by endohydrolases in
germinated barley grain. The enzymes could also play an important role
during cell wall turnover in elongating coleoptiles and in other
tissues during normal growth and development.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Peilin Xu for providing the aleurone cDNA library, Dr. Ken Chalmers for assistance with the genetic mapping, Dr. Neil Shirley for help with the protein analyses, and Dr. Yoji Hayasaka and Kris Ferguson for elements of the mass spectrometric analyses. Professor Bruce Stone, Dr. Hugues Driguez, Dr. Ross De Gori, and Dr. Andrew Harvey provided valuable advice in various aspects of the project.
![]() |
FOOTNOTES |
---|
* This work was supported in part by grants from the Australian Research Council and the Grains Research and Development Corporation of Australia (to G. B. F.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AY029259 (ARA-I) and AY029260 (XYL).
Recipient of a postgraduate scholarship from the Grains Research
and Development Corporation of Australia.
§ To whom correspondence should be addressed. Tel.: 61-8-8303-7296; Fax: 61-8-8303-7109; E-mail: geoff.fincher@adelaide.edu.au.
Published, JBC Papers in Press, December 2, 2002, DOI 10.1074/jbc.M210627200
2 S. Khan, GenBank accession number AC009243, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
4NPA, 4'-nitrophenyl
-L-arabinofuranoside;
ARA-I,
-L-arabinofuranosidase;
HCA, hydrophobic cluster
analysis;
HPLC, high performance liquid chromatography;
MALDI-TOF, matrix-assisted laser desorption ionization time-of-flight;
4NPX, 4'-nitrophenyl
-D-xylopyranoside;
RACE, rapid
amplification of cDNA ends;
RFLP, restriction fragment length
polymorphism;
XYL,
-D-xylosidase.
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REFERENCES |
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