|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
J Biol Chem, Vol. 275, Issue 20, 15082-15089, May 19, 2000
From the Pea microsomes contain an Plant cells are surrounded by a primary cell wall that is
~10-100 times thicker than the plasma membrane. The cell wall has several important physiological roles including control of
morphogenesis, intercellular signaling, and pathogen interactions (1).
Plant primary walls are composed mainly of a polysaccharide framework that is deposited during growth, thus determining the cell shape and
size. The major components of the primary cell wall are cellulosic microfibrils, hemicellulosic polysaccharides, pectins, and
glycoproteins (2).
Xyloglucan (XG)1 is the major
hemicellulosic polysaccharide of dicotyledonous and non-graminaceous
monocotyledonous cell walls. Within the wall it is firmly bound to
cellulose microfibrils (2) via hydrogen bonds and thus may have a
stabilizing function. The structure of XG differs among species,
although all have a Although extensive progress has been made in understanding the
structure of cell wall components such as XG, the processes of their
biosynthesis and assembly are still poorly understood. Therefore, to
understand the process of cell wall biosynthesis, isolation of
glycosyltransferases and polysaccharide synthases becomes critical.
With the exception of cellulose and callose, which are synthesized at
the plasma membrane, wall polysaccharides are made in the Golgi (10),
and many glycosyltransferase activities associated with their
biosynthesis have been identified in plant microsomal fractions
(11-14). However, only two genes have been cloned at this time (15,
16).
In previous work from our cell wall group (15), procedures were
developed to purify the XG-fucosyltransferase (XG-FTase) from pea
microsomes. Two polypeptides (~63 and ~68 kDa) co-purified with the
activity, and both of the bands were subjected to tryptic digestion and
sequencing. Peptides from the ~68-kDa band showed high sequence
similarity to binding protein (BiP), a molecular chaperone. However,
amino acid sequences from the ~63-kDa polypeptide identified one
clone of undetermined function in the Arabidopsis expressed
sequence tag data base. The full-length Arabidopsis cDNA
clone was isolated and shown to encode an ~63-kDa polypeptide with
XG-FTase activity (15). The goal of the present study is to complement
and extend our earlier work by isolating a pea cDNA clone for
XG-FTase (PsFT1), analyzing the biochemical characteristics of the
purified pea enzyme, determining the substrate acceptor specificity,
and characterizing the product of this enzyme in more detail. We show
that the purified transferase is indeed XG-specific. When tamarind XG
was used as an acceptor, the fucosylated subunits were the nona- and
decasaccharides, -XXFG- and -XLFG-, respectively. These two subunits
are found in native pea cell wall XG and also in many other plant XGs.
The characteristics of PsFT1 are compared with other described fucosyltransferases.
Chemicals and Plant Material
GDP-[3H]Fuc (299.7 GBq/mmol) and
GDP-[14C]Fuc (10.1 GBq/mmol) were purchased from NEN Life
Science Products. GDP-hexanolamine-agarose was a gift from Prof. Gerald
Hart (The Johns Hopkins University, Baltimore). BioSep-SEC-S4000 was
purchased from Phenomenex (Torrance, CA). Tamarind seed xyloglucan and
Trichoderma sp. cellulase were from Megazyme International
(Ireland). For earlier studies, tamarind seed XG was isolated as
described previously (17). Nucleotides, nucleotide sugars, all the
protease inhibitors, detergents, and the amino acid-modifying reagents
were from Sigma. Nasturtium seed XG and pea XG were a gift from Prof.
Gordon Maclachlan (McGill University, Montreal, Canada).
Rhamnogalacturonan-I and -II were a gift from Dr. Alan Darvill (Complex
Carbohydrate Research Center, University of Georgia, Athens, GA).
Garden pea seeds (Pisum sativum L. var. Alaska) purchased
from Olds Seed Company (Madison, WI) were washed in 10% bleach for 10 min, rinsed thoroughly, soaked 6-8 h with aeration, and grown in moist
vermiculate in the dark for 7 days until third internodes reached 3-5
cm in length. Growing regions (2 cm just below the hook) were excised
and used as the source of enzyme.
Purification Procedures
The purification was performed as described in the previous work
(15) except that in this study we did not include the anion exchange
column in purification steps.
PCR Amplification and Molecular Cloning
The cloning of PsFT1 was accomplished in several
steps. The first round PCR was performed using a plasmid cDNA
library as template and degenerate primers designed from peptides P5
and P2 (Fig. 1A) (5'-GCIGAYGGITTYGAYGARAA-3' and
5'-ICCCCAIACRTTRTTIGTIGGRTG-3'), where I indicates inosine; R
indicates purines (G + A), and Y indicates pyrimidines (T + C). The
cDNA library was derived from apical hooks of auxin-treated
etiolated pea seedlings in vector pBluescript SK(+) and was kindly
provided by Dr. Hans Kende (Department of Energy Plant Research
Laboratory, Michigan State University).
A second round PCR was performed to amplify fragments covering the 5'-
and 3'-ends. Primers were designed from the first round PCR
products (5'-TTCTACTAGTTGACCCGGGAGT-3' and
5'-ACTCCCGGGTCAACTAGTAGAA-3') and the cDNA library vector
(5'-TCAGGAAACAGCTATGACCATG-3' and 5'-GTAATACGACTCACTATAGGGC-3').
To identify the 5'- and 3'-ends of the PsFT1 transcript, 5'-
and 3'-RACE were carried out using the 5'-RACE System, version 2.0 (Life Technologies, Inc.), according to the manufacturer's instructions. Poly(A)+ RNA (gift from Dr. Hans Kende,
Department of Energy Plant Research Laboratory, Michigan State
University) was isolated from apical hooks of etiolated pea seedlings
4 h after auxin treatment (indoleacetic acid, 100 µM), using the poly(A)TRACT mRNA isolation system
(Promega). For 5'-RACE, 5'-ACTCCCGGGTCAACTAGTAGAA-3' was the
primer for cDNA synthesis, and
5'-GGCTGACTGATACCTGCTGAGACAAGATTTTT-3' was the nested primer. For
3'-RACE, poly(dT) primer was used for cDNA synthesis, and
5'-CCAAAAAGCCTGGGCAGAAAT-3' was the nested primer.
Full-length cDNA was amplified using Pwo polymerase (Roche
Molecular Biochemicals), end-specific primers
(5'-AATGAATATGCTGATAAAGAGAGTC-3' and
5'-CTAATTGTCTACGAGCTTAAGGC-3'), and the same cDNA
library as template. The same reaction was also performed using 300 ng genomic DNA as template to confirm the obtained sequence.
All PCR products were recovered from low melting point agarose gel
after electrophoresis and cloned onto the pGEM-T Easy vector (Promega)
for sequencing. All alignments were carried out using the software
package DNASTAR version 4.0 (Madison, WI).
Enzyme Assays
High Molecular Weight Polysaccharide Acceptors--
The standard
fucosyltransferase assay was performed essentially as developed (14,
18) with minor modifications. The reaction mixture (50 µl) contained
enzyme (up to 10 µg of protein), polysaccharides (100 µg) as
acceptor, GDP-[3H]Fuc (~3.3 nM, ~30,000
dpm) with 50 µM cold GDP-Fuc as donor, dissolved in the
assay medium (40 mM Pipes-KOH, pH 6.8, 0.4 mM DTT, 0.16 M sucrose), and supplemented with 5 mM MgCl2. In some experiments
GDP-[14C]Fuc (~2 µM, ~60,000 dpm) was
used without adding cold GDP-Fuc. Reactions were conducted for 20-30
min at room temperature and halted with 1 ml of 67% ethanol. Washed
ethanol-insoluble products were dissolved in 200 µl of water, and the
incorporated label (dpm) was measured in a Beckman LS 5000 TD Counter,
after adding at least 2 volumes of liquid scintillation mixture (ICN).
Control reactions without XG, without protein, or with boiled protein typically resulted in 50-100 dpm/reaction.
Low Molecular Weight Acceptors--
When lactose (5 mM), N-acetyllactosamine (5 mM), or
tamarind XG oligosaccharides (0.2% w/v) were used as acceptors, the
reactions were conducted as above and terminated by adding 300 µl of
water and then 500 µl of Dowex 1-X80 resin (Cl Glycoprotein Biochemical Characterization of the Purified XG-FTase
Size Estimation of the Purified Enzyme--
The apparent
Mr of the enzyme subunit was determined by
SDS-PAGE under reducing conditions (20) and by gel filtration using a
BioCAD Perfusion Chromatography system (Perspective Biosystems) and a
BioSep-SEC-S4000 column (Phenomenex) as described earlier (15).
pH Variation and Effectors--
pH stability of the enzyme was
investigated by incubating the enzyme (1 µl, ~0.3 µg) with 3 µl
of 66 mM phosphate buffer at pH 3.5, 4.2, 5.2, 6.3, 6.8, 7.8, 8.4, or 9.2 for 30 min on ice. The fucosyltransferase reaction was
then performed for 20 min in the standard assay using a higher
concentration of Pipes, pH 6.2 (46 µl of 0.1 M), to
equilibrate the pH in the reaction medium. Reactions were terminated as
above. The pH optimum was determined by conducting the assay in 50 mM solution of the following buffers: phosphate buffer, pH
5.2, 6.3, 6.8, 7.8, or 8.4; acetate buffer, pH 5.2; Bis-Tris-HCl, pH
6.8; Hepes-NaOH, pH 6.9; Tris-HCl, pH 7.5 or 8.0, containing all the
ingredients of the standard assay medium.
To assess whether divalent cations are required for activity,
MgCl2, MnCl2, or CaCl2 were
included at concentrations up to 10 mM in
fucosyltransferase assays lacking metal ions using ~2 µM GDP-[14C]Fuc (~60,000 dpm) as
described above. The effects of a chelator (EDTA) and DTT were
similarly tested.
Kinetic Studies--
Determination of kinetic parameters was
carried out using a 4 × 3 matrix of GDP-Fuc and tamarind XG in a
standard assay conditions (in absence of MgCl2). XG-FTase
velocity was measured as a function of tamarind XG (0.5, 1, or 4 mg/ml)
at fixed GDP-Fuc concentrations (12.5, 25, 50, or 100 µM). Reaction mixtures contained a fixed amount of
GDP-[3H]Fuc (~3.3 nM, ~30,000 dpm) and
enough purified enzyme to have an accurate initial reaction rate in 20 min incubation time. Data were evaluated by double-reciprocal plots
according to the Lineweaver-Burk method, and Km and
Vmax values were calculated from duplicate data
points (Fig. 3).
Amino Acid Modification--
Purified enzyme was subjected to
amino acid group-specific modification. The following reagents were
used: N-ethylmaleimide (NEM), diethylpyrocarbonate (DEPC),
pyridoxal 5'-phosphate (PLP), and N-bromosuccinimide (NBS).
Modifying reagents were used according to the methods described by
Britten and Bird (21). Standard assays were conducted as described
above in the presence of up to 5 mM of each modifying
reagent using GDP-[3H]Fuc. The reactions were halted by
adding 67% ethanol and washed as described above before assessing
label incorporation. Controls were conducted in the absence of these
modifying reagents.
Substrate Specificity--
The substrate acceptor specificity
was investigated by including alternative acceptors only or by
conducting competitive inhibition experiments. The reactions used
GDP-[14C]Fuc (~2 µM). The polysaccharides
used were tamarind XG, nasturtium XG, pea cell wall XG, and
rhamnogalacturonan-I and -II. Tamarind XG oligosaccharide subunits
monomers (7-9 degrees of polymerization) and trimers (21-27 degrees
of polymerization) were prepared by partial or complete digestion with
Trichoderma cellulase and purification on Bio-Gel columns.
Known acceptors for glycoprotein fucosyltransferases (lactose,
N-acetyllactosamine, and glycoprotein
Donor substrate specificity was investigated in a competition assay
using tamarind XG as acceptor. Nucleotides, nucleotide sugars, and
monosaccharides were tested by inclusion in the standard fucosyltransferase assay at various concentrations (up to 10 mM).
Identification of the Fucosylated Subunits of the XG
Fucosylated subunits in [14C]tamarind XG were
determined as described earlier (9, 14), with minor modifications.
After digestion by Trichoderma cellulase (0.05%, w/v) and
fractionation by gel filtration on a Bio-Gel P2 (Bio-Rad) column, the
samples (~10 µg of oligosaccharides, ~100-200 dpm/µg) were
further separated by HPAEC using a CarboPac PA-100 column (Dionex
Corp.) with pulsed amperometric detection (PAD). Fractions of 0.65 ml
were collected, neutralized, and analyzed for 14C
distribution. The positions of labeled oligosaccharides were compared
with those of authentic XG subunits as described (9, 14) and identified
according to the suggested nomenclature (3). Labeled products formed by
pea microsomes were also analyzed.
Molecular Cloning of PsFT1 and Sequence Analysis
Following purification, tryptic peptides from each purified
protein were subjected to limited amino acid sequencing. Peptide sequence information derived from this process has been published previously (15), and Fig. 1A
depicts the location of the six peptides derived from the 63-kDa
polypeptide (P1 to P6).
Molecular Cloning of PsFT1--
As a first step, a 726-base pair
internal fragment of the pea XG-FTase was PCR-amplified. The nature of
this product was confirmed by its 61.6% identity with AtFT1
and the presence of a region encoding previously sequenced peptide P1
(Fig. 1A). Primers designed from the middle of this
fragment, together with vector-specific primers were used to amplify
the cDNA fragments toward the 5'- and 3'-ends of this gene. This
generated a combined cDNA sequence encompassing the entire first
round PCR product. It encoded all six peptide sequences previously
obtained from the purified enzyme (15) with one amino acid change in
peptide P1 (Fig. 1A), which may be a sequencing error. To
ensure that we had the entire cDNA sequence of this gene, we
further performed 5'- and 3'-RACE experiments. The full-length
PsFT1 cDNA was then obtained by PCR from the cDNA library using primers specific to the 5'- and 3'-ends of the open reading frame and was consistent with the corresponding genomic clone
of PsFT1 (data not shown). The deduced amino acid sequence of PsFT1 is shown in Fig. 1A.
Characterization of the Deduced Amino Acid Sequence--
The
deduced translation product is 565 amino acid residues long, has a
predicted molecular mass of 64 kDa, and has a theoretical pI of 5.91 as
determined by DNASTAR (Madison, WI). A transmembrane domain was
predicted by TMHMM from residues 39 to 61 (Fig. 1), consistent with the
observation that Golgi-localized glycosyltransferases are type II
integral membrane proteins. An overall 62.3% identity was found
between PsFT1 and AtFT1. The highest similarity was observed at the
COOH terminus, postulated in other characterized fucosyltransferases to
contain the catalytic domain (22).
Analysis of Conserved Motifs--
Although the two XG-FTase
proteins have very low overall sequence similarity with the known
fucosyltransferases (less than 20% identity), three conserved motifs
were found. By using the PATTERNFIND program, we searched for motifs
(
Biochemical Characterization and Molecular Cloning of an
-1,2-Fucosyltransferase That Catalyzes the Last Step of Cell
Wall Xyloglucan Biosynthesis in Pea*
,§,¶,
,**,,, and**§§
Department of Energy Plant Research
Laboratory, the Cell and Molecular Biology Program, the
** Department of Botany and Plant Pathology, and the
Department of Biochemistry, Michigan State
University, East Lansing, Michigan 48824
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-fucosyltransferase
that incorporates fucose from GDP-fucose into xyloglucan, adding it
preferentially to the 2-O-position of the galactosyl
residue closest to the reducing end of the repeating subunit. This
enzyme was solubilized with detergent and purified by affinity
chromatography on GDP-hexanolamine-agarose followed by gel filtration.
By utilizing peptide sequences obtained from the purified enzyme, a
cDNA clone was isolated that encodes a 565-amino acid protein with
a predicted molecular mass of 64 kDa and shows 62.3% identity to its
Arabidopsis homolog. The purified transferase migrates at
~63 kDa by SDS-polyacrylamide gel electrophoresis but elutes from the
gel filtration column as an active protein of higher molecular weight
(~250 kDa), indicating that the active form is an oligomer. The
enzyme is specific for xyloglucan and is inhibited by xyloglucan
oligosaccharides and by the by-product GDP. The enzyme has a neutral pH
optimum and does not require divalent ions. Kinetic analysis indicates
that GDP-fucose and xyloglucan associate with the enzyme in a random
order. N-Ethylmaleimide, a cysteine-specific modifying
reagent, had little effect on activity, although several other amino
acid-modifying reagents strongly inhibited activity.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-1,4-glucan backbone with xylosyl side chains
attached to the 6-O-position of the glucosyl residues in a
regular pattern of three substituted glucosyl residues followed by one
unsubstituted glucose at the reducing end. L-Fucose
commonly terminates these side chains and is usually present in a
nonasaccharide subunit (Glc4Xyl3GalFuc), designated by agreement (3) as XXFG, where X is isoprimeverose (xylosyl--1,2-glucose) and F is the tetrasaccharide GlcXylGalFuc. Pea cell wall XG is thought to be constituted almost completely of
alternating -XXXG- (Glc4Xyl3) and -XXFG-
subunits (4), but recent study (5) has also detected the presence of
small amounts of other subunits including two octasaccharides and the
decasaccharide -XLFG-
(Glc4Xyl3Gal2Fuc). This subunit
(-XLFG-) has also been detected in sycamore cell XG (6),
Arabidopsis and rapeseed (7), apple fruit (8), and
developing nasturtium fruits (9). An unadecasaccharide, -XFFG-
(Glc4Xyl3Gal2Fuc2), has
additionally been observed in sycamore cell XG (6).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
form) as
described earlier (19). After centrifugation, the supernatants were
saved, and the resin was washed with 2 × 1 ml of water. All the
supernatants were combined, and the incorporated label was assessed as above.
1-Acid as Acceptor--
When
glycoprotein 1-acid (200 µg per reaction) was used as
acceptor, the reactions were halted by addition of 1 ml of 0.1% bovine
serum albumin and 1 ml of an acid mixture of 1 M HCl
containing 2% (w/v) phosphotungstic acid as described earlier (19).
After centrifugation, the pellets were washed three times with 1 ml of
acid mixture and 1 ml of methanol and then resuspended in 400 µl of
water. The labeled product was counted as above.
1-acid)
were also tested.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (57K):
[in a new window]
Fig. 1.
Predicted amino acid sequences of XG-FTase
from pea (PsFT1) and Arabidopsis
(AtFT1). A, sequence alignment
using the ClustalW algorithm. The six tryptic peptides obtained from
micro-sequencing of purified pea enzyme (P1 to
P6) are underlined. Black shading
represents conserved amino acids found in AtFT1 and PsFT1, and
gray shading represents similar amino acids. The conserved
motif found in all known -1,2- and -1,6-fucosyltransferases is
boxed. B, hydropathic plot of PsFT1 according to
the method of Kyte and Doolittle (31). The predicted transmembrane
domain is indicated in B by TMD.
-2/6-motif I, -2-motif II, -2-motif III, -6-motif II, and
-6-motif III) previously identified by Oriol et al. (23)
that are present in all known -1,2- and -1,6-fucosyltransferases.
The -2/6-motif I, present in all these fucosyltransferases, was
found in both PsFT1 and AtFT1. An additional motif was also found in
PsFT1 and AtFT1 at the proper spacing for motifs III observed in all
known fucosyltransferases. Both the -2-motif III and -6-motif III
are present in PsFT1 and AtFT1 (Fig. 2).
Interestingly, the -2-motif III and -6-motif III were previously
thought to be specific to known -1,2- and -1,6-fucosyltransferases, respectively (23). The -2-motif II and
-6-motif II were not found in PsFT1 and AtFT1. Fig. 2 summarizes the
relative locations of the conserved motifs, their spacing, and their
comparison to the consensus motifs of -1,2- and
-1,6-fucosyltransferases originating from other species.
View larger version (27K):
[in a new window]
Fig. 2.
Schematic illustration of peptide motifs
shared by XG-FTases and known -1,2- and
-1,6-fucosyltransferases. Conserved motifs in
AtFT1 and PsFT1 (boxed) are aligned (ClustalW) with the
corresponding motifs from some -1,2-fucosyltransferases from human
(H and Se) and bacteria (Yersinia
enterocolitica), from some -1,6-fucosyltransferases from human
(h6-FT), Caenorhabditis elegans-D (C. eleg. D), and two bacterial proteins (NodZ-B and
NodZ-C) from Rhizobium species
(Bradyrhizobium japonicum B and Rhizobium sp. C,
respectively). The invariant or highly related residues are shown in
white letters on black shading and other
conserved amino acids are in black letters on gray
shading. The positions of the first residues of each motif found
in the PsFT1 sequence are indicated on the bottom. The
GenBankTM accession numbers of the fucosyltransferases are
indicated on the right.
Biochemical Characterization of Purified XG-FTase
The enzyme recovered from the gel filtration column was used for subsequent biochemical characterization studies.
pH Stability and Optimum-- We investigated the effect of pH on the stability and activity of the purified enzyme, which was outside of a membrane environment and surrounded only by detergent. Purified pea XG-FTase was remarkably stable at extreme pH levels. Even after 30 min treatment with acidic (pH 3.5) or alkaline (pH 9.2) conditions, the enzyme retained over 80% of its activity (data not shown). This is consistent with stability during carbonate wash at pH 10.5. The fucosyltransferase was active between pH 5 and 8 with an optimum activity between pH 6 and 7 (data not shown).
Complete inhibition of the pea XG-FTase activity was observed in acetate buffer at pH 5.2, whereas a decrease of ~40% of the activity was observed with phosphate buffer at pH 5.2 (data not shown). Hepes, Pipes, and Bis-Tris at pH 6.8 did not interfere with the activity.
Effects of Divalent Cations and DTT-- Divalent metal ions are required for many proteins that bind nucleotide sugars. To investigate the metal ion dependence of XG-FTase, activity was measured in the presence and absence of various divalent cations. Activity was detected in the absence of cations or in the presence of EDTA (up to 10 mM) (data not shown), indicating that divalent cations are not required for its activity. However, the presence of 2 mM MgCl2 or CaCl2 in the assay medium enhanced the activity up to 30%. In contrast, MnCl2 had no effect at concentrations lower than 2 mM but at higher concentrations inhibited the enzyme (up to 60%) (data not shown). Adding DTT (up to 0.5 mM) enhanced the activity up to 2-fold and did not prevent the inhibitory effect of MnCl2 (data not shown).
Kinetic Analysis of XG-FTase--
The rate of XG-FTase activity
was determined at various concentrations of XG or GDP-Fuc,
respectively, and assessed by Lineweaver-Burk plots. A reciprocal plot
of 1/v against 1/S at a range of GDP-Fuc and
tamarind XG concentrations gave a series of straight lines that
intersect on the 
1/S axis (Fig.
3). This pattern indicates that GDP-Fuc
and tamarind XG have separate binding sites and that the binding of one
substrate has no consequence for binding of the other. Thus, we
concluded that XG-FTase interacted with the substrates in a random
order.
|
), 1 mg/ml (
), or 4 mg/ml (
). B, kinetic
values for XG-FTase at four concentrations of GDP-Fuc as follows: 12.5 µM (
), 25 µM (
), 50 µM
(
), or 100 µM (
). GDP-[3H]Fuc was
used, and data are expressed as picomoles of fucose incorporated per
min.
The Km for GDP-Fuc was ~30 µM and Vmax was ~150 mM/h/mg protein. Earlier data (24) for the unpurified XG-FTase activity indicated a Km of 72 µM, with a Vmax of ~0.8 mM/h/mg protein for GDP-Fuc. These results showed that the purified XG-FTase was more active with higher affinity. For tamarind XG, the Km was ~0.46 mg/ml (~0.40 µM) and Vmax ~200 mM/h/mg protein. The very low Km for XG may be because of a higher number of potential acceptor sites throughout the tamarind XG polymer, which increases its interaction with the enzyme.
Chemical Modification of Amino Acids-- To gain some insight about the structure-function relationship in XG-FTase, catalytically essential amino acid residue types were identified. The sulfhydryl reagent NEM, a cysteine-specific modifying agent, has been widely used in discriminating between glycoprotein fucosyltransferase isotypes (25). Our results indicated that NEM had little effect on the XG-FTase activity, with 5 mM giving ~40% inhibition of activity (Table I). However, N-bromosuccinimide (NBS), DEPC, and PLP reagents (which react with tryptophan, histidine and lysine residues, respectively) decreased the activity about 90-99% when applied at levels up to 5 mM (Table I). Preincubation of the XG-FTase with GDP-Fuc did not prevent inactivation by NBS treatment (data not shown). More studies using site-directed mutagenesis are needed to determine the locations and the roles of these essential residues.
|
Substrate Specificity-- To study the acceptor specificity of the purified XG-FTase, various substrates (polysaccharides, oligosaccharides, and glycoprotein) were first tested as potential acceptors (Table II). In a second experiment, some of these potential acceptors were tested for their ability to inhibit competitively the transfer of fucose onto tamarind XG (Fig. 4).
|
|
Table II and Fig. 4 show that rhamnogalacturonan-I and -II, both known
to contain terminal -fucosyl residues (26), were not substrates for
the purified pea XG-FTase. In addition, no incorporation was obtained
with 1-acid glycoprotein, despite the fact that it is a
good acceptor for animal glycoprotein -1,2- and
-1,3-fucosyltransferases. Tamarind XG is the best acceptor for the
fucosyltransfer reaction, and although nasturtium XG is also an
acceptor, it shows ~50% less incorporation than tamarind XG. One
likely explanation is that nasturtium XG has less (~2 mol %)
octasaccharide XXLG than tamarind XG (~31 mol %), and the position
of galactose in XXLG is preferentially fucosylated by the enzyme (14)
(see below). When pea XG was used, very little [14C]Fuc
incorporation (13% of the control) was obtained, probably because pea
XG is already almost completely fucosylated.
Tamarind XG monomers and trimers are comparatively poor acceptors (Table II, low Mr acceptors) but are able to compete with tamarind XG polymer. Strong inhibition was exhibited by the trimers (Fig. 4). Rhamnogalacturonan-II, lactose, and N-acetyllactosamine could not inhibit the fucosyltransferase reaction (Fig. 4). All these data indicate that the purified enzyme is indeed a xyloglucan-specific fucosyltransferase without any contaminant fucosyltransferase activities for glycoproteins or pectins.
To examine the specificity of the GDP-fucose-binding site, nucleotides
and nucleotide sugars were used as inhibitors of the fucosyltransferase
reaction. The result indicated that guanosine nucleotides provided
strong inhibition of XG-FTase in the following order: GMP 
GDP > GTP (data not shown). This can be explained as end product
competition. GDP-fucose showed the same inhibitory effect as GDP or
GMP, but no inhibition was obtained with fucose alone, suggesting that
the enzyme principally recognizes the nucleotide portion of the
GDP-fucose. However, the enzyme is specific for GDP-Fuc because GDP-Man
and GDP-Glc did not inhibit the reaction as efficiently as GDP-Fuc
(data not shown). Adenosine nucleotides did not provide significant inhibition.
Product Analysis, Identification of Fucosylated Subunits of the XG
Our previous work (15) demonstrated that the purified pea enzyme
transfers fucose to position 2 of tamarind XG side chain galactosyl
residues. However, we wanted to determine which of the three different
subunits found in tamarind XG, namely -XXLG-, -XLXG-, and -XLLG- (14),
was the preferred site of fucosylation for pea enzyme. Moreover, we
sought to determine which of the two galactosyl residues was
fucosylated when -XLLG- was used. To address these questions, tamarind
XG was fucosylated by the purified pea enzyme and was digested with
Trichoderma cellulase before fractionation on Bio-Gel P2
columns. The radiolabeled fragments eluted in a peak that overlapped
with the higher Mr side of the XG
oligosaccharides peak as expected (data not shown). The products were
further fractionated by HPAEC on a CarboPac PA-100 column (Fig.
5), and positions of labeled fragments
were compared with those of authentic XG oligosaccharides as described
previously (9, 14). Double peaks (one large and one very small) were observed, both with the radiolabeled products and with the marker oligosaccharides. The cause of the double peaks is not certain, but a
possible explanation could be that the minor peak represents the
oxidized form of the oligosaccharides at the reducing carbon at
position 1 of glucose to form gluconic acid residue. These charged
forms would elute slightly later than the parent oligosaccharides. Since all the authentic oligosaccharides behave in the same way, our
assignments regarding their structures are correct.
|
Radiolabel was recovered only in fractions where XXFG and XLFG eluted, at a dpm ratio of approximately 1.5:1, respectively (Fig. 5). There was no sign of products fucosylated closer to the non-reducing end of the subunits, e.g. XFXG or XFLG. We would expect these subunits to elute differently from one another, just as the octasaccharides do (Fig. 5). There was no indication of any labeled XFFG in these products, which would have been expected to elute from this column at a different locus from nona- and decasaccharide, as it does after high pressure liquid chromatography on a Dynamax-60A NH2 column (6). We conclude that the purified pea XG fucosyltransferase is specific for fucosylation of galactosyl residues nearest the reducing end of XG subunits.
This experiment was repeated using a crude enzyme preparation from
microsomal membranes in an effort to identify any isozymes present that
could generate different fucosylated subunits, namely XFXG, XFLG, or
XFFG. The data in Fig. 5 show that the crude enzyme exhibits the same
profile as the purified enzyme, but the dpm ratio between XXFG and XLFG
has changed to 3:1. This suggests that, in tamarind XG, the
membrane-bound form of the enzyme specifically fucosylates the -XXLG-
subunit over -XLLG-. This could be due to a regulatory factor present
in the microsomes or a modification of -XLFG- by either -fucosidase
or -galactosidase, which could increase the amount of XXFG over XLFG.
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
The XG-specific fucosyltransferase was first detected in vitro in microsomal membrane preparations from pea epicotyl tissue (27). Fucosylation did not require concurrent addition of UDP-Glc, UDP-Xyl, or UDP-Gal and was evidently due to transfer of fucose to endogenous nascent XG. Microsomal membranes incorporated several times more [14C]Fuc from GDP-[14C]Fuc into added tamarind XG than into endogenous XG (18). Isopycnic centrifugation provided evidence that the XG-FTase is present in both a dense Golgi fraction and in vesicles from the trans-Golgi network (28). Later studies using antibodies to the fucosylated region of XG determined that the fucose-containing product is observed in the trans-Golgi cisternae and in the trans-Golgi network (10). Taken together, these studies provide evidence that fucosylation occurs after XG backbone biosynthesis has been completed.
This XG-specific fucosyltransferase from pea epicotyl microsomes was purified as described earlier (15). Analysis of the purification data indicates that XG-FTase is present at very low levels in the cells (less than 0.01% of total protein).
Gel filtration and SDS-PAGE data indicated that the native form of
XG-FTase exists as an oligomer (~250 kDa), possibly including the BiP
chaperone protein (~68 kDa polypeptide) as part of the complex. The
biological relationship between the two polypeptides is still unknown.
One possibility is that the chaperone protein BiP may be involved
directly or indirectly in XG biosynthesis. A similar dependence has
been recently shown for the cell wall -1,6-glucan synthesis in
Saccharomyces cerevisiae (29), where -1,6-glucan
biosynthesis depends partly on the molecular chaperone BiP/Kar2p. The
other possibility is that the interaction between XG-FTase and BiP is
simply an artifact resulting from the carbonate treatment of the
microsomes. Nonetheless, many -1,2-fucosyltransferases for
glycoprotein studied to date have an oligomeric form with a molecular
mass of ~150 kDa, but SDS-PAGE estimations of molecular mass are
~50 kDa (22). It has been postulated that oligomerization might
function as a signal for retention in the Golgi, avoiding transport to
the plasma membrane (30).
As expected, PsFT1 shared a high degree of amino acid sequence
similarity with its Arabidopsis homolog (AtFT1). Comparison of both sequences provided further support for the conclusion that the
cloned pea cDNA encoded a XG-FTase enzyme. The Kyte-Doolittle hydrophilicity plot (31) suggested the presence of a 23-amino acid
transmembrane domain (TMD) at the NH2 terminus, indicating that PsFT1 is a type II membrane-bound protein with the active site
located in the lumen, like all other fucosyltransferases described to
date (with the exception of the bacterial fucosyltransferases that lack
the TMD). The conclusion that PsFT1 is a type II membrane protein is
also supported by the observation that activity was greatly stimulated
upon the addition of detergents (i.e. BRIJ 58) to sealed
microsomal membrane vesicles without protein
solubilization.2 Computer
analysis indicated that PsFT1 and AtFT1 shared three conserved motifs
with known -1,2- and -1,6-fucosyltransferases from vertebrate,
invertebrate, and bacteria. One very recent study (23) using Fasta
search and hydrophobic cluster analysis showed that -1,2- and
-1,6-fucosyltransferases have two common motifs (motif I and motif
II) in addition to one motif (motif III) specific to each group.
Interestingly, for the Arabidopsis and pea XG-FTases the
motif I is present, but the motif III in these XG-FTases contains elements from both known -2-motif III and -6-motif III. This favors a common genetic origin for these three groups of transferases.
Biochemical characterization revealed that PsFT1 enzyme was stable over a wide pH range (3-9), required no divalent cations for full activity, and had a random order interaction with its substrates as has been observed for other fucosyltransferases (32-35).
Several amino acid modification effects were observed. A characteristic
that clearly distinguishes different subtypes of fucosyltransferases is
the inactivation by a low amount of NEM due to an essential cysteine
residue involved in GDP-Fuc binding (35). NEM had little or no effect
on the pea XG-FTase activity. However, reagents that modify tryptophan,
histidine, and lysine residues (NBS, DEPC, and PLP, respectively)
significantly reduced the activity. Histidine is often found at active
sites of enzymes where its imidazole group charge can change depending
on local environment and thus catalyze the cleavage of bonds. PLP is
known to inhibit effectively enzymes that bind phosphorylated
substrates via a Schiff base with an amino group. For example, a
GDP-fucose-protectable pyridoxal-5'-P-modifiable Lys residue has been
shown to be present in the -1,3-fucosyltransferase from human small
cell lung carcinoma NCI-H69 cells (36). A more recent study showed that
a Lys residue located within an -1,3-fucosyltransferases motif is
required for activity but not substrate binding (37). It has been shown
also that this basic residue is involved in a hydrogen bond between
donor and acceptor hydroxyl groups during the catalysis (38).
Interestingly, two conserved Lys residues are found located within this
segment in PsFT1 and AtFT1 enzymes. Since divalent cations were not
required for the activity of PsFT1, it is possible that the Lys
residues may serve the same function as the cations by neutralizing the
negative charge of the phosphate on GDP-Fuc to enable it to associate
with the enzyme. The purified pea XG-FTase was extremely sensitive to
NBS, demonstrating an essential tryptophan residue that could not be
protected by GDP-Fuc. This suggests that the tryptophan residue is
located at a site distinct from the sugar nucleotide-binding site.
PsFT1 has 32 lysine, 12 histidine, and 8 tryptophan residues in its
amino acid sequence. Most of them are conserved with its
Arabidopsis homolog AtFT1 and are located at the COOH terminus.
The purified XG-FTase was indeed specific for XG, being unable to
utilize acceptors derived from either plant cell wall polysaccharides or glycoprotein carbohydrate moieties. It was also unable to fucosylate free tamarind XG oligosaccharides. However, competitive inhibition experiments indicate a higher affinity of the enzyme for the trimers than for the monomers. This result supports the recently proposed model
for the mode of action of pea XG-FTase activity (14) wherein the enzyme
binds to a trimer in galactosylated nascent chains and fucosylates the
octasaccharide (-XXLG-) nearest the reducing end of the chain.
Repetition of this action results in a chain with repeating
hepta:nonasaccharide subunits (-XXXG·XXFG-). It will be of interest
to investigate the minimal structure recognition of the acceptor with
this enzyme in the future. Also, it is of interest to know the
structural features that dictate acceptor substrate specificity. It has
been shown that only a few amino acids found within a
"hypervariable" region at the NH2 terminus of human
-1,3- and -1,4-fucosyltransferases determine the substrate specificity for these enzymes (39).
The products formed by the purified enzyme were identified as XXFG and XLFG following cellulase digestion of the 14C-fucosylated tamarind XG. This result was predicted because both of these subunits are found in XG (4-9). In pea XG, however, the predominant natural fucosylated subunit is XXFG; XLFG is a minor component (5). Thus, pea XG-FTase generated the same two fucosylated subunits in vitro as in vivo, but the ratio of their yields is different from that found in natural pea cell wall XG. This difference occurs because the products recovered in this experiment must have arisen by 14C-fucosyl transfer to the subunits -XXLG- and -XLLG-, the major subunits in tamarind XG, with a ratio of 3:5 (8, 14). It is interesting to point out that no products such as XFXG or XFLG were observed. Moreover, the existence of these oligosaccharides has not yet been demonstrated to date in any plant cell wall XG. The fact that the fucosylation occurs only at the galactosyl residues closest to the unbranched glucose in the XG indicates the existence of a recognition system that can discriminate between the two galactosyl residues in the same subunit (e.g. -XLLG-). This discrimination prevents the fucosylation of the second galactose in pea. A crude enzyme preparation from microsomal membranes produced mostly XXFG, showing that the pea tissues from growing regions probably have only one XG-FTase that makes this linkage. However, sycamore cells have fucose on both galactosyl residues in the same subunit (-XFFG-) (6). In this species it will be interesting to determine whether a second FTase exists or whether they have a single enzyme that is able to add fucose to both galactosyl residues.
The availability of the cDNA clones and information about the
biochemical properties of XG-FTase will allow future studies aimed at
understanding how this enzyme activity is regulated as cell wall
synthesis progresses during growth. It will also be important to
identify other enzymes involved in XG biosynthesis so that the entire
process can be analyzed.
| |
ACKNOWLEDGEMENTS |
|---|
We thank Prof. Hans Kende and Dr. Qin Du (Department of Energy Plant Research Laboratory, Michigan State University) for offering pea cDNA library and mRNA; Prof. Gerald Hart (The Johns Hopkins University, Baltimore) for the GDP-agarose column; Prof. Gordon Maclachlan (McGill University, Montreal, Canada) for pea cell wall XG and nasturtium seed XG; Dr. Alan Darvill (Complex Carbohydrate Research Center, University of Georgia) for rhamnogalacturonan-I and -II; Dr. Anton Sanderfoot for continued advice; and all members of the Keegstra and Raikhel laboratories for their support.
| |
FOOTNOTES |
|---|
* This work was supported by the United States Department of Energy.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/EMBL Data Bank with accession number(s) AF223643.
§ Present address: Washington University School of Medicine, Dept. of Molecular Microbiology, Campus Box 8230, 660 South Euclid Ave., St. Louis, MO 63110-1093.
¶ Present address: Seattle Biomedical Research Institute, 4 Nickerson St., Suite 200, Seattle, WA 98109.
§§ To whom correspondence should be addressed. Tel.: 517-353-7874; Fax: 517-353-9168; E-mail: keegstra@msu.edu.
Published, JBC Papers in Press, March 9, 2000, DOI 10.1074.jbc.M000677200
2 A. Faik, unpublished data.
| |
ABBREVIATIONS |
|---|
The abbreviations used are: XG, xyloglucan; XG-FTase, xyloglucan fucosyltransferase; PsFT1, Pisum sativum fucosyltransferase 1; AtFT1, Arabidopsis thaliana fucosyltransferase 1; PAGE, polyacrylamide gel electrophoresis; DTT, dithiothreitol; PCR, polymerase chain reaction; RACE, rapid amplification of cDNA ends; HPAEC, high pH anion exchange chromatography; PAD, pulsed amperometric detection; GDP-hexanolamine, P'-(6-amino-1-hexyl)-P2-(5'-guanosine)pyrophosphate; DEPC, diethylpyrocarbonate; NEM, N-ethylmaleimide; PLP, pyridoxal 5'-phosphate; NBS, N-bromosuccinimide; Bis-Tris, 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol; Pipes, 1,4-piperazinediethanesulfonic acid; TMD, transmembrane domain.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
| 1. | McNeil, M., Darvill, A. G., Fry, S. C., and Albersheim, P. (1984) Annu. Rev. Biochem. 53, 625-663 |
| 2. | Carpita, N., and Gibeaut, D. (1993) Plant J. 3, 1-30 |
| 3. | Fry, S. C., York, W. S., Albersheim, P., Darvill, A. G., Hayashi, T., Joseleau, J.-P., Kato, Y., Lorences, E. P., Maclachlan, G. A., McNeil, M., Mort, A. J., Reid, J. S. G., Seitz, H., Selvendran, R. R., Shibear, R. R., Voragen, A. G. J., and White, A. R. (1993) Physiol. Plant. 89, 1-3 |
| 4. | Hayashi, T., and Maclachlan, G. (1984) Plant Physiol. 75, 596-604 |
| 5. | Guillén, R., York, W. S., Pauly, M., An, J., Impallomeni, G., Albersheim, P., and Darvill, A. G. (1995) Carbohydr. Res. 277, 291-311 |
| 6. | Hisamatsu, M., Impallomeni, G., York, W. S., Albersheim, P., and Darvill, A. G. (1991) Carbohydr. Res. 211, 117-129 |
| 7. | Zablackis, E., Huang, J., Müller, B., Darvill, A. G., and Albersheim, P. (1995) Plant Physiol. 107, 1129-1138 |
| 8. | Vincken, J.-P., Beldman, G., Niessen, W. M. A., and Voragen, A. G. J. (1996) Carbohydr. Polym. 29, 75-85 |
| 9. | Desveaux, D., Faik, A., and Maclachlan, G. (1998) Plant Physiol. 118, 885-894 |
| 10. | Zhang, G. F., and Staehelin, L. A. (1992) Plant Physiol. 99, 1070-1083 |
| 11. | Bolwell, G. P., and Northcote, D. H. (1984) Planta 162, 139-146 |
| 12. | Waldron, K. W., and Brett, C. T. (1985) in Biochemistry of Plant Cell Wall (Brett, C. T. , and Hillman, J. R., eds) , pp. 79-97, Cambridge University Press, New York |
| 13. | Doong, R. L., Liljebjelke, K., Fralish, G., Kumar, A., and Mohnen, D. (1995) Plant Physiol. 109, 141-152 |
| 14. | Faik, A., Chileshe, C., Sterling, J., and Maclachlan, G. (1997) Plant Physiol. 114, 245-254 |
| 15. | Perrin, R. M., DeRocher, A. E., Bar-Peled, M., Zeng, W., Norambuena, L., Orellana, A., Raikhel, N. V., and Keegstra, K. (1999) Science 284, 1976-1979 |
| 16. | Edwards, M. E., Dickson, C. A., Chengappa, S., Sidebottom, C., Gidley, M. J., and Reid, G. (1999) Plant J. 19, 691-697 |
| 17. | Edwards, M. E., Dea, I. C. M., Bulpin, P. V., and Reid, J. S. G. (1985) Planta 163, 133-140 |
| 18. | Farkas, V., and Maclachlan, G. A. (1988) Arch. Biochem. Biophys. 264, 48-53 |
| 19. | Hokke, C. H., Neeleman, A. P., Koeleman, C. A. M., and van den Eijnden, D. H. (1998) Glycobiology 8, 393-406 |
| 20. | Laemmli, U. K. (1970) Nature 227, 680-685 |
| 21. | Britten, C. J., and Bird, M. I. (1997) Biochim. Biophys. Acta 1334, 57-64 |
| 22. | Kleene, R., and Berger, E. G. (1993) Biochim. Biophys. Acta 1154, 283-325 |
| 23. | Oriol, R., Mollicone, R., Cailleau, A., Balanzino, L., and Breton, C. (1999) Glycobiology 9, 323-334 |
| 24. | Hanna, R., Brummell, D. A., Camirand, A., Hensel, A., Russel, E., and Maclachlan, G. (1991) Arch. Biochem. Biophys. 290, 7-13 |
| 25. | de Vries, T., Srnka, C. A., Palcic, M. M., Swiedler, S. J., van den Eijnden, D. H., and Macher, B. A. (1995) J. Biol. Chem. 270, 8712-8722 |
| 26. | Darvill, A. G., McNeil, M., and Albersheim, P. (1978) Plant Physiol. 62, 418-422 |
| 27. | Camirand, A., and Maclachlan, G. A. (1986) Plant Physiol. 82, 379-383 |
| 28. | Brummell, D., Camirand, A., and Maclachlan, G. (1990) J. Cell Sci. 96, 705-710 |
| 29. | Simons, J. F., Ebersold, M., and Helenius, A. (1998) EMBO J. 17, 396-405 |
| 30. | Nilson, T., Hoe, M. H., Slusarewicz, P., Rabouille, C., Waton, R., Hunte, F., Watzele, G., Berger, E. G., and Warren, G. (1994) EMBO J. 13, 562-574 |
| 31. | Kyte, J., and Doolittle, R. F. (1982) J. Mol. Biol. 157, 105-132 |
| 32. | Beyer, T. A., and Hill, R. L. (1980) J. Biol. Chem. 255, 5373-5379 |
| 33. | Murray, B. W., Takayama, S., Schultz, J., and Wong, C. H. (1996) Biochemistry 35, 11183-11195 |
| 34. | Macher, B. A., Holmes, E. H., Swiedler, S. J., Stults, C. L. M., and Srnka, C. A. (1991) Glycobiology 1, 577-584 |
| 35. | Prieels, J.-P., Monnom, D., Dolmans, M., Beyer, T. A., and Hill, R. L. (1981) J. Biol. Chem. 256, 10456-10463 |
| 36. | Holmes, E. H. (1992) Arch. Biochem. Biophys. 296, 562-568 |
| 37. | Sherwood, A. L., Nguyen, A. T., Whitaker, J. M., Macher, B. A., Stroud, M. R., and Holmes, E. (1998) J. Biol. Chem. 273, 25256-2526 |
| 38. | Hindsgaul, O., Kaur, K. J., Srivastava, G., Blaszczyk-Thurin, M., Crawley, S. C., Heerze, L. D., and Palcic, M. M. (1991) J. Biol. Chem. 266, 17858-17862 |
| 39. | Legault, D. J., Kelly, R. J., Natsuka, Y., and Lowe, J. (1995) J. Biol. Chem. 270, 20987-20996 |
Copyright © 2000 by The American Society for Biochemistry and Molecular Biology, Inc.
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  O. Zabotina, E. Malm, G. Drakakaki, V. Bulone, and N. Raikhel Identification and Preliminary Characterization of a New Chemical Affecting Glucosyltransferase Activities Involved in Plant Cell Wall Biosynthesis Mol Plant, November 1, 2008; 1(6): 977 - 989. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. M. Cavalier, O. Lerouxel, L. Neumetzler, K. Yamauchi, A. Reinecke, G. Freshour, O. A. Zabotina, M. G. Hahn, I. Burgert, M. Pauly, et al. Disrupting Two Arabidopsis thaliana Xylosyltransferase Genes Results in Plants Deficient in Xyloglucan, a Major Primary Cell Wall Component PLANT CELL, June 1, 2008; 20(6): 1519 - 1537. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. Ma, J. L. Simala-Grant, and D. E. Taylor Fucosylation in prokaryotes and eukaryotes Glycobiology, December 1, 2006; 16(12): 158R - 184R. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. M. Cavalier and K. Keegstra Two Xyloglucan Xylosyltransferases Catalyze the Addition of Multiple Xylosyl Residues to Cellohexaose J. Biol. Chem., November 10, 2006; 281(45): 34197 - 34207. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Geisler-Lee, M. Geisler, P. M. Coutinho, B. Segerman, N. Nishikubo, J. Takahashi, H. Aspeborg, S. Djerbi, E. Master, S. Andersson-Gunneras, et al. Poplar Carbohydrate-Active Enzymes. Gene Identification and Expression Analyses Plant Physiology, March 1, 2006; 140(3): 946 - 962. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Shao, H. Zheng, Y. Hu, D. Liu, J.-C. Jang, H. Ma, and H. Huang The GAOLAOZHUANGREN1 Gene Encodes a Putative Glycosyltransferase that is Critical for Normal Development and Carbohydrate Metabolism Plant Cell Physiol., October 15, 2004; 45(10): 1453 - 1460. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
X. Li, I. Cordero, J. Caplan, M. Molhoj, and W.-D. Reiter Molecular Analysis of 10 Coding Regions from Arabidopsis That Are Homologous to the MUR3 Xyloglucan Galactosyltransferase Plant Physiology, March 1, 2004; 134(3): 940 - 950. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Madson, C. Dunand, X. Li, R. Verma, G. F. Vanzin, J. Caplan, D. A. Shoue, N. C. Carpita, and W.-D. Reiter The MUR3 Gene of Arabidopsis Encodes a Xyloglucan Galactosyltransferase That Is Evolutionarily Related to Animal Exostosins PLANT CELL, July 1, 2003; 15(7): 1662 - 1670. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. M. Perrin, Z. Jia, T. A. Wagner, M. A. O'Neill, R. Sarria, W. S. York, N. V. Raikhel, and K. Keegstra Analysis of Xyloglucan Fucosylation in Arabidopsis Plant Physiology, June 1, 2003; 132(2): 768 - 778. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. Freshour, C. P. Bonin, W.-D. Reiter, P. Albersheim, A. G. Darvill, and M. G. Hahn Distribution of Fucose-Containing Xyloglucans in Cell Walls of the mur1 Mutant of Arabidopsis Plant Physiology, April 1, 2003; 131(4): 1602 - 1612. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. Joly, R. Leonard, A. Maftah, and C. Riou-Khamlichi {alpha}4-Fucosyltransferase is regulated during flower development: increases in activity are targeted to pollen maturation and pollen tube elongation J. Exp. Bot., June 1, 2002; 53(373): 1429 - 1436. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Faik, N. J. Price, N. V. Raikhel, and K. Keegstra An Arabidopsis gene encoding an alpha -xylosyltransferase involved in xyloglucan biosynthesis PNAS, May 28, 2002; 99(11): 7797 - 7802. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 V. Chazalet, K. Uehara, R. A. Geremia, and C. Breton Identification of Essential Amino Acids in the Azorhizobium caulinodans Fucosyltransferase NodZ J. Bacteriol., December 15, 2001; 183(24): 7067 - 7075. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. R. Hicks, C. M. Hironaka, D. Dauvillee, R. P. Funke, C. D'Hulst, S. Waffenschmidt, and S. G. Ball When Simpler Is Better. Unicellular Green Algae for Discovering New Genes and Functions in Carbohydrate Metabolism Plant Physiology, December 1, 2001; 127(4): 1334 - 1338. [Full Text] [PDF]
|
|
|
|
|
|
R. Sarria, T. A. Wagner, M. A. O'Neill, A. Faik, C. G. Wilkerson, K. Keegstra, and N. V. Raikhel Characterization of a Family of Arabidopsis Genes Related to Xyloglucan Fucosyltransferase1 Plant Physiology, December 1, 2001; 127(4): 1595 - 1606. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Vitale Uncovering Secretory Secrets: Inhibition of Endoplasmic Reticulum (ER) Glucosidases Suggests a Critical Role for ER Quality Control in Plant Growth and Development PLANT CELL, June 1, 2001; 13(6): 1260 - 1262. [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| All ASBMB Journals | Molecular and Cellular Proteomics |
| Journal of Lipid Research | ASBMB Today |