The Plant Journal (1999) 20(6), 629±639
Molecular domains of the cellulose/xyloglucan network in
the cell walls of higher plants
Markus Pauly², Peter Albersheim, Alan Darvill and
William S. York*
Complex Carbohydrate Research Center and Department
of Biochemistry and Molecular Biology, University of
Georgia, 220 Riverbend Road, Athens, GA 30602-4712,
USA
Summary
Cellulose and xyloglucan (XG) assemble to form the
cellulose/XG network, which is considered to be the
dominant load-bearing structure in the growing cell
walls of non-graminaceous land plants. We have
extended the most commonly accepted model for the
macromolecular organization of XG in this network,
based on the structural and quantitative analysis of
three distinct XG fractions that can be differentially
extracted from the cell walls isolated from etiolated pea
stems. Approximately 8% of the dry weight of these cell
walls consists of XG that can be solubilized by
treatment of the walls with a XG-speci®c endoglucanase
(XEG). This material corresponds to an enzymesusceptible XG domain, proposed to form the cross-links
between cellulose micro®brils. Another 10% of the cell
wall consists of XG that can be solubilized by
concentrated KOH after XEG treatment. This material
constitutes another XG domain, proposed to be closely
associated with the surface of the cellulose micro®brils.
An additional 3% of the cell wall consists of XG that can
be solubilized only when the XEG- and KOH-treated cell
walls are treated with cellulase. This material constitutes
a third XG domain, proposed to be entrapped within or
between cellulose micro®brils. Analysis of the three
fractions indicates that metabolism is essentially limited
to the enzyme-susceptible domain. These results
support
the
hypothesis
that
enzyme-catalyzed
modi®cation of XG cross-links in the cellulose/XG
network is required for the growth and development of
the primary plant cell wall, and demonstrate that the
structural consequences of these metabolic events can
be analyzed in detail.
Received 16 August 1998; revised 12 October 1998; accepted 21 October
1998.
*For correspondence (fax +1 706 542 4412; e-mail [email protected]).
²
Present address: Plant Biochemistry Laboratory, Department of Plant
Biology, The Royal Veterinary and Agricultural University, 40 Thorvaldsensvej, DK-1871 Frederiksberg C, Denmark.
ã 1999 Blackwell Science Ltd
Introduction
All higher plant cells are encased in a cell wall, which
de®nes the cell's shape and thereby contributes to the
structural integrity of the entire plant. Growing cells are
surrounded by a metabolically active primary cell wall,
which is capable of expanding (Kerr and Bailey, 1934). The
primary cell wall of all higher plants consists of crystalline
cellulose micro®brils, embedded in a hydrated (65% water;
Fry, 1988), amorphous matrix of hemicelluloses, pectins,
and glycoproteins (Darvill et al., 1980) that form a network
around the cell. According to current cell-wall models, the
cellulosic framework is interconnected by hemicellulosic
polysaccharides, such as xyloglucan (XG) or arabinoxylan,
forming a cellulose/hemicellulose network (Carpita and
Gibeaut, 1993). Co-existing with the cellulose/hemicellulose network is another network that consists of the pectic
polysaccharides homogalacturonan, rhamnogalacturonan
I and rhamnogalacturonan II. In addition, primary cell walls
often contain structural proteins such as the hydroxyproline-rich glycoprotein extensin.
The structural rigidity and strength of the wall is thought
to depend on the integrity of the cellulose/hemicellulose
network. Enzyme-catalyzed modi®cation of the hemicellulosic component of this network is considered to be
essential for wall expansion during cell growth (Talbott
and Ray, 1992). A more complete knowledge of the
structure, macromolecular organization, and metabolism
of this network is necessary to understand the mechanisms leading to plant cell elongation and the biological
regulation of this process.
The structural components of the cellulose/hemicellulose network are well known. Cellulose micro®brils consist
of non-covalently associated, linear chains of b-1,4-linked
D-glucopyranosyl residues. XG is the major hemicellulosic
component in dicotyledonous and non-graminaceous
monocotyledonous plants. Like cellulose, XG has a b-1,4
linked glucospyranosyl backbone, which is branched, with
many of its b-D-glucosyl residues bearing an a-D-xylosyl
residue at C6 (Figure 1). The xylosyl residues can be further
substituted to form oligomeric side chains containing
galactosyl, fucosyl, and/or arabinosyl residues. The structure and molecular distribution of these XG side chains
varies in different plant tissues and species. The chemical
structures of these side chains have been rigorously
established by analysis of xyloglucan oligosaccharides
(XGOs) that are generated upon endoglucanse digestion of
the polymer. However, relatively little is known about the
macromolecular organization of XG in the cell wall.
629
630 Markus Pauly et al.
Perhaps the most important feature of XGs is their
characteristic capacity to form strong, non-covalent associations with cellulose (Valent and Albersheim, 1974;
Hayashi, 1989; Hayashi et al., 1994). XGs bind to cellulose
in vitro in a pH-dependent manner (Hayashi et al., 1987),
suggesting that the formation of hydrogen bonds is
involved in the association of these polymers.
Concentrated alkali (e.g. 24% KOH) is required to extract
XG from the cellulose/XG complex (Hayashi, 1989; Hayashi
and Maclachlan, 1984). Antibody studies (Vian et al., 1992)
suggest that XGs are associated with the surface of
cellulose micro®brils in muro. Cellulose micro®brils
cross-linked by putative XG `tethers' have been visualized
by electron microscopy of rotary shadowed replicas of
rapidly frozen, deep-etched cell-wall specimens (McCann
et al., 1990; McCann et al., 1992; Itoh and Ogawa, 1997) and
of arti®cially assembled bacterial cellulose/XG composites
(Whitney et al., 1995). XG cross-links were also suggested
by studies in which cell-wall epitopes recognized by an
XG-speci®c antibody were observed predominantly in the
spaces between micro®brils (Baba et al., 1994).
This paper describes a model for the cellulose/XG
network based on the observation that differentially
extractable xyloglucan fractions are structurally distinct.
Three different macromolecular domains are proposed for
the XG in this network. The data presented here suggest
that the metabolism of XG associated with cell expansion
occurs predominantly in the most accessible of the three
domains.
Results
Quantitative solubilization of xyloglucans from pea-stem
cell walls by a sequential extraction procedure
Figure 1. Two xyloglucan-derived oligosaccharides (XGOs).
Endoglucanases such as cellulase or XEG hydrolyze the unbranched b-Dglucosyl residues of the XG backbone, to generate XGOs such as those
shown here. XGO structures described in the text and in Table 4 are
speci®ed using a standard nomenclature (Fry et al., 1993), in which an
uppercase letter indicates the precise side-chain substitution pattern of
each b-D-glucosyl residue in the oligosaccharide. The letter G represents
an unbranched b-D-glucosyl residue, and the letter X represents a b-Dglucosyl residue with an a-D-xylosyl substituent at O6, etc. Thus, the
sequences XXXG and XLFG unambiguously de®ne the two structures
shown.
The macromolecular organization of XG in the cell walls of
etiolated pea (Pisum sativum L.) stems was investigated.
This tissue was chosen because: (i) the structural features
of XG in this tissue are well established (Hayashi et al.,
1984; Guillen et al., 1995); (ii) suf®cient quantities of this
tissue can be easily obtained; and (iii) metabolic turnover
of XG in this tissue has been studied previously (Hayashi
and Maclachlan, 1986; Talbott and Ray, 1992). XG was
solubilized from the pea-stem cell walls using a sequential
extraction procedure (see Experimental procedures).
Destarched cell walls were partially depectinated by
treatment with a puri®ed endopolygalacturonase (EPG) in
combination with pectin methylesterase (PME). This
procedure increases the amount of XG that can be
solubilized by the subsequent steps (data not shown),
con®rming a previous observation (Bauer et al., 1973) and
suggesting that removal of pectin increases the enzyme
accessibility of XG.
The depectinated cell walls were treated with an XGspeci®c endoglucanase (XEG), which selectively solubilizes XGOs (Pauly et al., 1999). Under the conditions used,
this treatment solubilized approximately 8.2% of the wall
material (Table 1). XEG was removed from the soluble
extract by passing it through an anion-exchange resin and
the salts were removed by size-exclusion chromatography
(SEC) on Sephadex G-10. The sugar composition of the
desalted fraction con®rmed that only XGOs are present in
this fraction (Table 1). The use of additional XEG, extended
digestion times, or multiple XEG treatments did not
signi®cantly increase the amount of wall material solubilized by this method (data not shown). Colorimetric
analysis of the extract indicated that XEG solubilized
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
Cellulose/xyloglucan network
XG, the KOH extract contained acidic components of the
cell wall, including pectic polymers and glucuronoarabinoxylan, which were removed by passing the extract
through a column of Q-Sepharose. (Prior to Q-Sepharose
chromatography, the desalted extract was treated with
XEG to generate XGOs.) The monosaccharide composition
of the neutral ¯owthrough (i.e. the XGO-enriched fraction,
Table 1) was consistent with the presence of XG (glucose,
xylose, galactose, and fucose), but arabinose was also
abundant (42.4%), probably due to the presence of a
neutral arabinoxylan or an arabinan. A small amount of
mannose in the extract suggested that it also contained a
mannan. Quanti®cation of the XG present in this fraction
by the anthrone method was not signi®cantly compromised by the presence of the arabinose or xylose because
the anthrone assay is insensitive to these pentoses.
However, the anthrone assay is sensitive to mannose,
and the sugar-composition data in Table 1 were used to
estimate the colorimetric response of mannose in the KOH
extract. After taking this `mannose factor' into account, the
anthrone assay indicated that XG in the KOH-solubilized
material comprised 10.3% of the initial mass of the
partially depectinated cell walls (Table 1).
The amount of XG remaining in the cell walls after XEG
and KOH treatment was determined by analysis of the
monosaccharides released by Saeman hydrolysis of the
residue. Saeman hydrolysis involves the use of concentrated H2SO4 (see Experimental procedures) to depolymerize all of the polysaccharide components of the wall,
including microcrystalline cellulose (Selvendran et al.,
1979). The resulting monosaccharides are not signi®cantly
degraded (Selvendran et al., 1979) and can be quantitated
by GLC analysis of their alditol acetate derivatives
(Table 2). This analysis was also performed on cell walls
that had not been treated with XEG and KOH. Under the
8.0% of the cell walls as XGOs, in good agreement with the
gravimetrically determined decrease in the insoluble cellwall material (8.2%) observed upon XEG treatment.
Additional XG was solubilized when the XEG-treated
walls were extracted with concentrated alkali (24% KOH).
Approximately one-third of the initial mass of the partially
depectinated wall material was solubilized by this treatment (Table 1), but further base treatment did not release a
signi®cant amount of additional material. In addition to
Table 1. Yields and sugar compositions of XGO-enriched
fractionsa,b obtained by sequential extraction of pea-stem CWM
with XEG, KOH, and cellulase
Parameter
XEG
KOH
Cellulase
8.2
8.0
33.8
10.3
27.0
3.9
ndg
3.3
nd
34.3
nd
12.8
49.6
nd
0.9
42.4
23.8
3.3
4.7
24.9
nd
2.1
nd
32.1
nd
9.5
56.3
c
Yield (wt%)
Amount solubilizedd
Recovered as XGOse
Composition (mol%)f
Rhamnose
Fucose
Arabinose
Xylose
Mannose
Galactose
Glucose
631
a
XGO-enriched fractions were prepared as described in
Experimental procedures.
b
Data are the average of two extraction experiments.
c
Yields are wt% relative to the initial mass of the partially
depectinated pea-stem cell wall.
d
Determined by mass reduction of the residual CWM.
e
Determined by the anthrone assay.
f
Sugar compositions are normalized mol% of each XGOenriched fraction.
g
Not detected.
Table 2. Sugar composition (mass %) of Saeman hydrolysates of cell-wall material
Sugars
Partially depectinated
cell walls (untreated)
Cell walls treated
with XEG and KOHa
Cell walls treated
with XEG, KOH, and cellulasea
Totalb
Rhamnose
Fucose
Arabinose
Xylose
Mannose
Galactose
Glucose
100
0.1c
0.3
11.5
6.6
4.8
2.7
74.0
58
ndd
nd
0.9
1.6
2.0
0.3
53.2
31
nd
nd
nd
0.1
0.2
0.8
29.9
a
Cell walls were sequentially extracted.
The data are normalized by reference to the total mass of the untreated cell walls, which is set to 100%. As indicated in Table 1, 58%
of the cell wall remained insoluble after sequential treatment with XEG and KOH, and 31% remained insoluble after subsequent
treatment with cellulase.
c
Average of two extraction experiments.
d
Not detected.
b
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
632 Markus Pauly et al.
conditions described above, 58% of the cell-wall mass
remained after sequential extraction with XEG and KOH.
Therefore, the mass percentages of the monosaccharides
(Table 2) present in the residual cell walls were normalized
to give a total of 58%. Glucose is the most abundant sugar
detected in both treated and untreated cell walls. Most of
this glucose probably originated from the hydrolysis of
cellulose, as cellulose accounts for up to 30% of the dry
weight of the cell walls of higher plants (Fry, 1988). The
monosaccharide-composition data shown in Table 2 indicate that sequential XEG and KOH treatment of the cell
walls solubilized signi®cant amounts of several cell-wallmatrix polysaccharides (XG, xylans, mannans, and pectic
polysaccharides). Most of the cellulose remained in the
insoluble residue (hereafter called the XEG/KOH residue).
The XEG/KOH residue was treated with cellulase, which
solubilized nearly all of the remaining matrix polysaccharides but less than half of the insoluble cellulose.
It is dif®cult to determine accurately the amount of XG
present in the insoluble cell-wall fractions. The presence of
cellulose in these fractions makes quanti®cation of glucose
a poor indicator of their XG content. However, quantitative
analysis of xylose can be used to estimate the XG content
of the XEG/KOH residue because xylose is present in all
XGOs. The XEG/KOH residue contains xylose, which could
be due to the presence of xylan and/or XG. However,
xylans are more readily solubilized by KOH than are XGs
(York et al., 1985). Therefore, most of the xylose in the
XEG/KOH residue is probably derived from XG that is not
solubilized by this treatment. Analysis (described below) of
the oligosaccharides released by cellulase treatment of the
XEG/KOH residue is consistent with this conclusion.
Retreatment of the XEG/KOH residue with XEG was used
in an attempt to solubilize more XGOs, as the KOH
treatment might have exposed some of the XG that had
previously been inaccessible to XEG. However, no additional wall material was released by the second XEG
treatment (data not shown). The ability of a commercially
available cellulase to solubilize additional XGOs was also
tested. Cellulase treatment solubilized an additional 27% of
the partially depectinated walls (Table 1). Two carbohydrate-rich fractions were obtained when the cellulasesolubilized material was desalted by SEC (data not shown)
(Pauly et al., 1999). The sugar composition and elution
volume of the very low-molecular-weight fraction indicated that it contained cellobiose and glucose, the
products expected upon enzymatic degradation of cellulose. The sugar composition (Table 1) and elution volume
of the second fraction indicated that XGOs were its main
constituents, and suggested that it may also have
contained a small amount of larger cello-oligosaccharides
that were co-eluted with the XGOs. Quantitative analysis of
the second fraction by the anthrone method showed that
XG released by this cellulase treatment consitituted an
additional 3.9% of the cell wall (Table 1). The insoluble
residue obtained after sequential treatment with XEG,
KOH, and cellulase comprised 31% of the mass of the
depectinated cell wall. Therefore, the mass percentages of
the monosaccharides (Table 2) present in this a-cellulose
fraction were normalized to give a total of 31%. As
expected, glucose is the most abundant sugar present
from this fraction. Xylose constituted only 0.2% of this
fraction, indicating that xylose-containing wall polysaccharides, including XG, are almost quantitatively solubilized by the sequential extraction procedure.
These results show that the sequential extraction
procedure, employing XEG, KOH (24%), and ®nally
cellulase, solubilized three distinct domains of the XG
present in the cell walls. Here, domain is not de®ned as a
structural entity within a molecule, but rather as an
`environmental' domain based on its extractability.
However, the results do not establish whether individual
XG molecules occupy a single domain or span more than
one domain.
Connectivities between the different solubilized XG
domains
Additional insight into the physical relationships among
the various XG domains was obtained by analysis of XG
that was extracted by KOH with no prior XEG treatment. In
this case, KOH-solubilized XG accounted for 18.4% of cell
walls (data not shown). Thus, KOH alone solubilizes
approximately the same amount of XG as sequential
extraction with XEG and KOH (18.3%, Table 1), suggesting
that XEG-extractable XGs can also be solubilized with
KOH. Furthermore, KOH treatment of the wall does not
break glycosidic bonds in the XG backbone, releasing XG
as a polysaccharide.
These results suggested the following experiments to
determine whether the XEG- and KOH-extractable domains are covalently connected. The molecular mass
distributions of KOH-solubilized XG obtained with or
without prior XEG treatment of the cell wall material
(CWM) were determined by SEC on Superose 12 (Figure 2).
Fractions were collected and assayed using the anthrone
assay, which detects hexoses, and iodine staining, which
selectively detects XGs with a molecular weight greater
than approximately 20 kDa. The molecular weights of the
KOH-solubilized XGs were estimated from their SEC
elution volumes, using dextrans and tamarind XGOs as
standards.
Two carbohydrate peaks dominate the chromatogram of
the KOH extract prepared without prior treatment of the
CWM with XEG (Figure 2, top panel). The iodine- and
anthrone-positive peak indicated the presence of an XG
with an average molecular weight of 100 kDa. The smaller
anthrone-positive (iodine-negative) peak indicated the
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
Cellulose/xyloglucan network
633
arabinose content (data not shown), indicating the presence of an arabinan. Thus, prior treatment with XEG
decreased the molecular weight of the KOH-extracted XGs
from approximately 100 to 30 kDa, suggesting that individual XG molecules span two domains, of which one can be
solubilized by XEG while the other can be extracted only
upon treatment with KOH. The results are also consistent
with the hypothesis that the XEG-accessible XG domain is
interdispersed within the KOH-extractable XG domain.
One intriguing possibility is that, on average, an approximately 28 kDa stretch at each end of a typical 100 kDa XG
molecule is closely associated with cellulose (and thus
resistant to XEG), and that the intervening approximately
44 kDa is susceptible to XEG attack. This hypothetical
arrangement is consistent with both the SEC data (Figure
2) and the data in Table 1, as the two `cellulose-bound'
ends of the polysaccharides and the intervening `crosslink' would represent 10.3 and 8.0%, respectively, of the
dry weight of the cell wall.
In vitro experiments with an XG/Avicel composite
Figure 2. The effects of prior XEG treatment on the Superose 12 pro®le of
XGs solubilized by KOH treatment of partially depectinated pea-stem cell
walls.
Fractions were collected and assayed for hexoses by the anthrone assay
(d) and for XG by iodine staining (s). Both assays involved measuring
the absorption at 620 nm. Dextrans (superscript a) and BEPS-XGOs
(superscript b) of known molecular weight (expressed as kDa) were used
to calibrate the column, as indicated by the arrows.
presence of a low-molecular-weight (~2 kDa) glycan. The
monosaccharide composition of the two peaks indicated
that their main components were XG and arabinan/
arabinogalactan, respectively (data not shown). Two peaks
were also observed in the chromatogram obtained by SEC
of the KOH extract prepared from XEG-treated cell walls
(Figure 2, lower panel). A very small amount of material
was eluted at the void volume of the column, indicating
the presence of a large polymer with a molecular weight of
more than 500 kDa. This polymer did not stain with iodine
and consisted mainly of xylose (data not shown), indicating it was a xylan rather than an XG. The bulk of the
material solubilized by KOH after XEG treatment was
eluted at a volume corresponding to an average molecular
weight of approximately 30 kDa. Iodine staining and
monosaccharide composition of this material indicated
that it consisted mainly of XG, but also had a high
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
An arti®cial aggregate, composed of XG and microcrystalline cellulose (Avicel), was also analyzed by the sequential
extraction method. This system is largely free of other cellwall components that could complicate analysis of the
composite. As the binding of XGs to cellulose in vitro does
not involve enzymes, the molecular structure of the XG,
including its chain length and molecular topology, is
unlikely to be modi®ed in the binding process. It is also
unlikely that XG would become entrapped within a
cellulose micro®bril or irreversibly entangled with cellulose chains upon binding to Avicel. Thus, the molecular
topology of this arti®cial system is likely to be much
simpler than that of a plant cell wall synthesized in vivo.
This relative simplicity makes it possible to directly
examine the interaction of XG molecules with the surface
of previously formed cellulose micro®brils. The results
presented here demonstrate that this interaction, in itself,
cannot lead to the complex topology of a real cell wall.
Nevertheless, analysis of the XG/Avicel system indicated
that association of XG with the cellulose surface gives rise
to two distinct, differentially extractable XG domains that
are analogous to the XEG- and KOH-extractable XG
domains in pea stem cell walls.
XG obtained from the extracellular polysaccharides of a
bean suspension culture (BEPS-XG) was bound to Avicel
in vitro as described in Experimental procedures. The XG/
Avicel composite contained approximately 1 mg BEPS-XG
per 15 mg Avicel, as estimated by colorimetric analysis of
XGs remaining in the supernatant after incubation with the
cellulose (data not shown). The XG/Avicel composite and
`naked' Avicel were each treated with buffer, XEG,
cellulase, or KOH under the same conditions used to
634 Markus Pauly et al.
Table 3. Glycosyl composition (mg) material extracted from Avicel (15 mg) or the XG/Avicel composite (1 mg BEPS-XG/15 mg Avicel)
Avicel
XG/Avicel
Sugars
Buffer
XEG
Cellulase
KOH
Buffer
XEG
Cellulase
KOH
Rhamnose
Fucose
Arabinose
Xylose
Mannose
Galactose
Glucose
Total
nda
0.1
0.2
0.5
nd
0.8
3.0
4.6
nd
0.1
0.3
0.7
nd
0.5
3.2
4.8
nd
0.2
2.6
2.5
5.1
1.2
215.7
227.3
nd
0.3
3.5
8.3
87.5
1.1
322.1
454.8
nd
0.3
0.2
4.3
nd
2.4
9.4
16.6
nd
4.3
0.8
41.6
0.7
23.8
85.5
156.7
nd
9.3
3.6
113.2
42.6
66.0
769.0
1003.7
nd
13.5
4.6
301.4
129.8
81.9
722.1
1253.3
a
Not detected.
Table 4. XGO-compositiona,b of fractions solubilized by sequential XEG, KOH, and cellulase treatment
XEG
Oligosaccharidec
Not acetylated
Mono-acetylated
Di-acetylated
KOH
Not acetylatedd
Cellulase
Not acetylatedd
XXG
GXXG
XXXG
XLXG
XXLG
XLLG
GXFG
XXFG
XLFG
12.7
0.9
39.4
1.6
2.0
0.4
ni
11.4
2.4
nie
ni
ni
nd
5.5
1.5
1.2
13.5
7.1
ni
ni
ni
nd
nd
nd
ni
0.4
nd
5.0
ndf
40.3
1.5
8.6
1.3
nd
32.2
11.1
0.3
nd
47.7
1.8
6.9
1.9
nd
34.2
7.3
a
See Fry et al. (1993) and Figure 1 for a description of the XGO nomenclature used here.
Normalized mol%, average of two extraction experiments.
c
See Figure 1 for nomenclature.
d
All acetyl substituents are hydrolyzed by KOH treatment.
e
Not identi®ed: no standards are available for these particular O-acetylated forms, as we have never observed them as constituents of
cell wall XGs.
f
Not detected.
b
extract the pea cell wall. Each treatment was considered
`complete', as repeated extraction with a given reagent did
not solubilize additional carbohydrate (data not shown).
The monosaccharide components of each of the solubilized fractions were quanti®ed by GLC of their alditol
acetate derivatives (Table 3).
Buffer extraction of either naked Avicel or the XG/
Avicel complex released a negligible amount of carbohydrate. XEG released a total of 156 mg of carbohydrate
from the XG/Avicel complex. Thus, after accounting for
the amount of carbohydrate (4.6 mg) released upon XEG
treatment of naked Avicel, these results indicate that
approximately 15% of the XG in the complex was
released by XEG.
Considerably more carbohydrate was released by cellulase treatment of the XG/Avicel complex than by XEG
treatment. Most of the extracted material was glucose
generated by enzymatic depolymerization of the Avicel
itself. As expected, cellulase treatment of naked Avicel also
solubilized a considerable amount of glucose-containing
polysaccharides. Cellulase solubilized only 2.5 mg of xylose
from naked Avicel but 113.2 mg of xylose from the Avicel/
XG complex. Considering that xylose represents approximately 30% of the mass of BEPS-XG, and that 1 mg of
BEPS-XG is bound in the XG/cellulose complex, XG
contributes approximately 300 mg of xylose to the composite. Hence, cellulase treatment of the complex solubilized
approximately 37% of the bound XG.
KOH solubilized a large amount of glucose from naked
Avicel but only minute quantities of other sugars (e.g.
mannose, xylose, and arabinose). However, KOH treatment of the XG/Avicel complex released 301.4 mg of xylose,
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
Cellulose/xyloglucan network
635
Figure 3. Model accounting for differential extractability of XG domains in the XG/cellulose network.
This model emphasizes the likely arrangement of the various XG domains in different micro-environments, but is not meant to quantitatively represent the
mass ratio of these domains. Cellulose micro®brils are indicated by the grey bars. The XG domain that can be solubilized with XEG (blue bars) is
proposed to be accessible to enzyme-catalyzed modi®cation in vivo. This domain may form loops, dead ends, or cross-links between cellulose micro®brils.
The XG domain that can be solubilized by KOH (red bars), with or without prior XEG treatment, is bound to the surface of the cellulose micro®brils. The
XG domain that remains insoluble after XEG/KOH extraction of the cell wall (light green bars and dots in the cross-sections of some of the cellulose
micro®brils) is trapped within or between cellulose micro®brils. Some of this XG domain may also reside on the surface of cellulose micro®brils,
remaining resistant to XEG due to its close association with the micro®bril, and resistant to KOH extraction due to its covalent attachment to entrapped
XG. Little or none of the XG that can be solubilized by XEG appears to be covalently attached to entrapped XG.
indicating that essentially all of the XG in the complex was
solubilized by this treatment.
XGO composition of the XG-enriched fractions obtained
from pea stems
The XGO compositions of the sequentially solubilized
XEG, KOH, and cellulase fractions were determined in
order to establish whether any structural feature of the XG
is correlated with its presence in a particular domain.
Solubilized XGOs were reductively aminated with pnitrobenzylhydroxy-amine (PNB, Pauly et al., 1996). The
resulting PNB-XGOs were separated by reversed-phase
chromatography and quanti®ed by integrating their UV
absorbance (Pauly, 1999) (Table 4). O-Acetylated XGOs
present in the XEG-solubilized fraction were separated and
quanti®ed by this method. However, base treatment of the
walls hydrolyzes any O-acetyl substituents that may be
present, and O-acetylated glycans were not present in the
fractions solubilized by KOH and subsequent cellulase
treatment. The overall compositions of the three fractions
are very similar, with XXXG and XXFG being the main
oligosaccharide subunits in all XG domains ± see Figure 1
and Fry et al. (1993) for a description of the XGO
nomenclature. However, only the XEG-accessible domain
contains the `xylose-de®cient' oligosaccharides GXXG and
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
GXFG. Furthermore, the amount of XXG, an oligosaccharide that may be formed by enzymatic trimming of the nonreducing end of the XG, is correlated to the accessibility of
the domain, and is almost undetectable in the cellulaseextracted domain.
Discussion
An extended model for the XG/cellulose network in pea
stems
Most current models of the primary cell wall include a
cellulose/XG network in which the XG cross-links the
cellulose micro®brils (Hayashi and Maclachlan, 1984).
These models predict that at least two XG domains exist
in the cell wall, with one domain bound directly to cellulose
micro®brils and a second domain forming the cross-links.
We extend this model for the macromolecular organization
of the cellulose/XG complex based on the analysis of peastem cell walls (Figure 3). The pea epicotyl tissues used in
this study are heterogeneous mixtures of cell types. Thus,
this analysis cannot fully describe changes that occur in the
XG/cellulose network of any speci®c cell. However, the
overall trends that were observed reveal the presence of
three XG domains that are likely to occur in the cell walls of
most non-solanaceous, dicotyledonous plants. In our
636 Markus Pauly et al.
extended model, XG that is extractable by XEG corresponds to a molecular domain that constitutes the XG
cross-links between cellulose micro®brils and any exposed
tails or loops of XG that extend away from the micro®bril
surface. The XEG-extractable domain is covalently attached
to the KOH-extractable XG domain, which is non-covalently
bound to the surface of cellulose micro®brils. Close
association of the KOH-extractable domain with the micro®bril surface makes it inaccessible to the XEG. KOH
treatment of cell walls that have not been pretreated with
XEG releases both the XEG- and KOH-extractable domains.
A third XG domain is entrapped within or between cellulose
micro®brils, where neither XEG or KOH can gain access.
The entrapped XG may reside in relatively non-crystalline
(amorphous) regions of the micro®bril. This entrapped XG
can be released by cellulase treatment, which depolymerizes amorphous regions of the cellulose and hydrolyzes
the unbranched glucosyl residues in the XG backbone. The
entrapped XG domain consists of molecules that are rarely,
if ever, contiguous with the XEG- and KOH-extractable XG
domains, as the XEG and KOH domains can be solubilized
in their entirety (by KOH) without breaking covalent bonds
(i.e. without XEG pretreatment) and without releasing any
of the entrapped XG. These data do not rule out the
possibility that some of the entrapped XG molecules may
extend onto the surface of cellulose micro®brils. Such
molecular extensions may not be accessible to XEG, due to
their close association with the micro®bril surface, and
would not be solubilized by KOH, due to their covalent
attachment to the entrapped XG domain.
Further support for this model comes from analysis of
an XG/cellulose composite produced by mixing the two
components in vitro. This system is simpler than the XG/
cellulose network in the cell wall because XG is bound
exclusively to the surface of the micro®brils, and none is
internalized (trapped) as may occur in vivo when cellulose
is synthesized in the presence of XG. XG that binds to the
surface of cellulose in vitro may form loops or cross-links
between micro®brils. XEG treatment of this arti®cial
system solubilized approximately 15% of the bound XG;
cellulase treatment solubilized approximately 34% of the
bound XG; and KOH treatment solubilized virtually 100%
of the bound XG (Table 3). Even multiple XEG treatments
solubilize only a small portion of the XG, consistent with
the hypothesis that XEG can attack only the XG domain
that constitutes XG cross-links and/or loops. XG was more
ef®ciently solubilized by treating the XG/Avicel system
with cellulase than by treating it with XEG, consistent with
the hypothesis that partial depolymerization of cellulose
exposes additional XG, making it accessible to enzyme.
Virtually all of the XG in the arti®cial XG/cellulose system
was solubilized by KOH treatment, supporting the hypothesis that the KOH-extractable domain is closely associated
with the surface of the cellulose micro®bril.
It is clear from the results presented here that a
signi®cant portion of the XG molecules in the cell wall
span two molecular domains that are selectively solubilized by sequential XEG and KOH treatment. These two
domains almost certainly correspond to different microenvironments within the cell wall. The `enzyme-accessible
domain', operationally de®ned by its extractability by XEG,
represents the putative XG cross-link. The `surface-bound
domain' can be solubilized by KOH but not by XEG (Figure
3). This model predicts that the enzyme-accessible domain
can be modi®ed in vivo by plant cell-wall enzymes,
including endoglucanases (EGs), xyloglucan endotransglycosylases (XETs), and exoglycosidases such as afucosidases (Augur et al., 1992) or b-galactosidases (Ross
et al., 1993). The model also predicts that surface-bound
domain cannot be modi®ed by enzymes such as EGs and
XETs that attack the glucosyl backbone of the XGs, but
might be modi®ed by exoglycosidases that attack XG side
chains extending away from the micro®bril. A third
`trapped domain', corresponding to XG that is not
solubilized by XEG and KOH, is likely to be completely
inaccessible to cell-wall enzymes in vivo. Although the
enzyme-accessible XG domain is the most likely to be
modi®ed during the assembly and expansion of the cell
wall, it is important to note that our results do not establish
whether a given XG molecule can evolve from one domain
to another during plant cell development.
The physical basis for segregating XG into separate
domains in the primary cell wall is poorly understood. This
process is likely to be affected by the primary structure of
the XG and any enzyme-catalyzed modi®cations that affect
its primary structure or topology. For example, it is
possible that the presence of O-acetyl substituents could
affect the rate and extent to which an XG molecule binds to
cellulose. However, the degree of O-acetylation in the
surface-bound domain cannot be readily determined
because solubilization of this domain requires strong
alkali. Nevertheless, cellulose-binding experiments using
native (O-acetylated) and de-O-acetylated BEPS-XG indicated that the presence of O-acetyl groups does not affect
the amount of XG that binds to cellulose in vitro (Pauly,
1999). Another possibility is that binding to cellulose may
be modulated by the local sequence of XGO subunits in
the polysaccharide. Our data do not allow us to evaluate
this hypothesis because, although the XGO subunit
composition was determined for each of the three XG
domains, the order of the various XGOs in the original
polysaccharide was not.
Most XG subunits have an even number of glucosyl
residues (e.g. XXXG, XXFG), leading to a regular topology
that facilitates binding to cellulose (Levy et al., 1991). It has
been proposed that the binding of XG to cellulose can be
disrupted by a `topological reversal' that occurs at sites
consisting of XG subunits (e.g. XXG) that have an odd
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
Cellulose/xyloglucan network
number of glucosyl residues. This proposal is consistent
with the observation that the occurrence of XG subunits
(such as XXG) having an odd number of residues decreases
as the XG domain is more tightly associated with the
cellulose micro®bril (Table 4). However, it is also possible
that this trend is a result of intimate contact with cellulose,
rather than its cause. That is, XG subunits such as XXG may
arise by sequential trimming of exposed non-reducing ends
of the polysaccharide, converting, for example, XXXG to
GXXG and then to XXG. Such enzyme-mediated transformations are most likely to occur in the enzyme-accessible
domain, consistent with the data presented in Table 4.
The model presented here does not represent a radical
departure from previously published hypotheses concerning the macromolecular structure of XG in the primary cell
wall. However, the results demonstrate that there are three
distinct XG domains in the primary walls of pea stem cells,
and that it is possible to study the individual characteristics
of each of these domains. De®ning the genesis and
metabolism of these domains and analyzing their conformational, topological, and dynamic properties are
extremely challenging problems, the solutions to which
will require the application of a broad range of genetic,
biochemical, and physical techniques.
Experimental procedures
Chemicals, reagents, substrates and enzymes
Buffer salts, acids, bases, and organic solvents were obtained
from J.T. Baker (Philipsburg, NJ, USA); other reagents were
purchased from Sigma Chemical Co. (St Louis, MO, USA).
XGO fragments of the soluble XG secreted by suspensioncultured bean cells (BEPS-XGOs) were prepared as described
(Wilder and Albersheim, 1973).
Endopolygalacturonase (EPG) from Aspergillus niger was
obtained from Dr Carl Bergmann, CCRC. Pectin methylesterase
(PME) and xyloglucan-speci®c endoglucanase (XEG) (Pauly et al.,
1999) from Aspergillus aculeatus was obtained from NovoNordisk (Copenhagen, Denmark). XEG was puri®ed as described
(Pauly et al., 1999).
Colorimetric assays
The total carbohydrate content of samples was quanti®ed using
the anthrone assay for hexoses as described (Dische, 1962). A
known amount of BEPS-XGOs was used as a standard when
determining the XG or XGO content of a sample. The XG content
of samples was estimated by an iodine-staining assay (Kooiman,
1960). Samples were dissolved in 100 ml water, Gram-stain (75 ml
aqueous KI, 6.6 g l±1, and I2, 3.8 g l±1) and sodium sulfate (500 ml
aqueous Na2SO4, 0.2 g ml±1) were added, and the solution was
incubated for 1 h. A 200 ml aliquot of the solution was transferred
to a microtiter plate and the absorbance at 620 nm was measured.
The uronic acid content (e.g. galacturonic acid from pectic
polysaccharides) of samples was quantitated using the mhydroxybiphenyl assay (Blumenkrantz and Asboe-Hansen, 1973).
ã Blackwell Science Ltd, The Plant Journal, (1999), 20, 629±639
637
Isolation of cell walls
Cell walls were prepared from whole stems of 9-day-old etiolated
pea plants (Guillen et al., 1995). The entire isolation procedure
was carried out at 4°C. Harvested tissue was suspended (1 g
FW ml±1) in potassium phosphate buffer (100 mM, pH 7.0) containing 5 mM Na2S2O5 as an antioxidant, and homogenized in a
Polytron (Brinkman, Westburg, NY, USA) at maximum speed for
5 min. The homogenized tissue was ®ltered through a triple layer
of nylon mesh (1 mm pore size) and washed three more times
with the same buffer. The solid residue was resuspended in the
same volume of 500 mM phosphate buffer (pH 7.0) containing
5 mM Na2S2O5, ®ltered through the nylon mesh and washed three
more times with the same buffer. The residue was then
resuspended in the same volume of 0.5% aqueous SDS containing 3 mM Na2S2O5, and stirred for 20 h. The suspension was
®ltered and the solid residue was washed with water and then
resuspended in a 1 : 1 mixture of chloroform and methanol,
homogenized for 3 min with the Polytron, and ®ltered. The
insoluble residue (CWM) was then washed with acetone and
dried under vacuum at 30°C.
To remove starch, the CWM was suspended (0.5 g DW ml±1) in
100 mM potassium phosphate, pH 7.0, containing antibiotic
(0.01% thimerosal) and a-amylase (EC 3.2.1.1, Type IIA from
Sigma, 2.5 mg g±1 FW). The suspension was stirred at room
temperature for 48 h, then ®ltered through four layers of nylon
mesh, washed three times with deionized water, washed with
acetone, and dried.
The CWM was partially depectinated using a combination of
EPG and pectin methylesterase PME. Dried CWM was suspended
(10 mg DW ml±1) in 100 mM sodium-acetate buffer (pH 5.2) containing 0.01% thimerosal. EPG (®ve units) and PME (®ve units)
were added and the suspension was incubated for 24 h at 37°C.
Solubilized material was then removed by ®ltration through a
nylon membrane (10 mm pore size; Micron Separations Inc.,
Westboro, MA, USA) in a polysulfone ®lter funnel (50 ml capacity;
Gelman Sciences, Ann Arbor, MI, USA). The partially depectinated CWM (retained on the ®lter) was then dried.
Solubilization of XGOs using XEG
XG was solubilized from cell walls using XEG (Pauly et al., 1999)
that had been puri®ed to remove glycosidases that might
otherwise degrade XGOs. CWM was suspended (100 mg DW) in
10 mM sodium acetate (pH 4.5, 10 ml) containing 0.02% thimerosal. XEG (10 units) was added and the suspension was incubated
for 24 h at 37°C. After the XEG digestion, the suspension was
®ltered through a nylon membrane (10 mm pore size) in a
polysulfone funnel to separate `XEG-XGOs' (®ltrate) from the
XEG-treated CWM.
Solubilization of XGs using 24% KOH
XEG-treated CWM was suspended (100 mg DW) in 24% (w/v)
aqueous KOH (10 ml) containing 0.1% NaBH4. The suspension
was stirred at room temperature for 24 h and then ®ltered through
a nylon membrane (10 mm pore size) in a polysulfone funnel. The
®ltrate was neutralized with glacial acetic acid and salts were
removed by dialysis [3500 MWCO tubing (Spectrum, Houston,
TX, USA)] versus deionized water (4°C). The retentate, containing
the KOH-solubilized XG, was buffered by adding 1 M sodium
acetate, pH 4.5, to a ®nal concentration of 10 mM. XGOs were
638 Markus Pauly et al.
generated by adding puri®ed XEG (®ve units) and incubating the
solution for 24 h at 37°C.
Solubilization of XGs using cellulase
A commercial cellulase (Megazyme, Bray, Ireland) was used to
solubilize XG from the KOH-extracted CWM. The insoluble
residue recovered after KOH-treatment was suspended (100 mg
DW) in 10 mM sodium acetate, pH 4.5 (10 ml), containing 0.02%
thimerosal. Cellulase (10 units) was added and the suspension
was incubated for 48 h at 37°C. After the cellulase digestion the
suspension was ®ltered through a nylon membrane (10 mm pore
size) in a polysulfone funnel.
Isolation of XGOs
Solubilized XGOs were subjected to anion-exchange chromatography on a Q-sepharose column (Sigma) to remove the enzymes
(cellulase or XEG) or pectic polysaccharides (cosolubilized by 24%
KOH). The column (5 ml in a 1.5 3 15 cm plastic tube) was
equilibrated with 10 mM imidazole/HCl pH 7.0, and the solubilized
XGOs were then applied in the same buffer. The column was
washed with two column volumes of the same buffer to elute the
neutral components (XGOs), which were desalted by SEC on a
Sephadex G-10 column (1.5 3 80 cm, Sigma). The G-10 column
was eluted with deionized water, and collected fractions were
assayed for salt content by conductivity and for carbohydrate
content using the anthrone assay. The resulting `XGO-enriched
fractions' were characterized as described below.
XGO-composition
The XGO composition of each XGO-enriched fraction was
determined by HPLC of PNB-XGOs prepared by reductive
amination with p-nitrobenzylhydroxylamine hydrochloride, as
described (Pauly et al., 1996). PNB-XGOs were separated by
HPLC on a Vydac 238TP54 reversed-phase C-18 column (Vydac,
Hesperia, CA,USA). The sample was eluted from the column with
a linear gradient from 7 to 12% aqueous acetonitrile (1 ml min±1)
over 40 min, followed by a linear gradient from 12 to 23% aqueous
acetonitrile over 20 min. PNB-XGOs were detected by monitoring
the UV absorbance (275 nm) of the eluant with a Beckman UV 163
variable wavelength detector.
Saeman hydrolysis
Saeman hydrolysis (Selvendran et al., 1979) was used to hydrolyze plant cell walls for monosaccharide composition analysis.
Plant cell walls (2±4 mg) were wet with 100 ml aqueous myoinositol (1 mg ml±1, internal standard) in a screw-capped borosilicate test tube. Concentrated H2SO4 (300 ml) was added, the test
tube was capped, and the suspension was left for 3 h at RT with
occasional vortexing. The suspension was then diluted with
6.6 ml of distilled water and heated for 2 h at 100°C. After cooling,
the solution was transferred to a 50 ml Falcon tube and titrated
with saturated barium hydroxide [~35±40 ml] to pH 6±7 to remove
sulfate ions, which precipitate as Ba(SO4). The hydrolysate was
centrifuged for 5 min at 2000 g and the supernatant ®ltered
through ®lter paper (Whatman, Fair®eld, NJ, USA) into a roundbottomed ¯ask. The pellet was resuspended in water (40 ml) by
vortexing and centrifuged again. The resulting supernatant was
®ltered as described above and combined with the ®rst super-
natant. The pooled solution was concentrated by rotary evaporation to approximately 5±6 ml. The monosaccharide composition
of the Saeman hydrolysate was determined by analyzing (see
below) the alditol acetate derivatives prepared from 500 ml of the
solution.
Monosacharide composition analysis
The monosaccharide compositions of the various CWM fractions
were determined by gas±liquid chromatography (GLC) of alditol
acetate derivatives (York et al., 1985). Monosaccharides were
prepared by Saeman hydrolysis (Selvendran et al., 1979) of the
insoluble residues recovered after various extractions, or by
hydrolysis of water-soluble oligo- and polysaccharides in aqueous tri¯uoroacetic acid. The internal standard (myo-inositol) was
added to the samples before hydrolysis. The resulting monosaccharides were reduced with sodium borohydride and Oacetylated. A standard sample containing a mixture of rhamnose,
fucose, arabinose, xylose, mannose, galactose, and glucose was
prepared and derivatized in parallel with the experimental
samples.
XG binding to cellulose
Cellulose (Avicel PH101, Fluka, Milwaukee, WI, USA) was
prewashed at least three times by suspending it (15 mg) in 15 ml
incubation buffer (50 mM sodium acetate, pH 4.5, room temperature), centrifuging at 2000 g for 5 min in a Marathon 6K table-top
centrifuge (VWR, McGaw Park, IL, USA), and decanting the
supernatant. The washed Avicel was resuspended in 1 ml of the
incubation buffer and BEPS-XG (1.5 mg) was added. The suspension was vigorously shaken in an incubator (180 r.p.m.) for 12 h at
room temperature. After this binding period, the bound and
unbound XG was separated by centrifuging the suspension
(2000 g for 5 min) and decanting the supernatant. The insoluble
Avicel±XG complex was washed twice with 500 ml incubation
buffer. The Avicel±XG complex and a control consisting of Avicel
without added XG were then treated with XEG, cellulase, or KOH,
under the conditions (described above) used to extract XG from
CWM. After each treatment the insoluble residue (XG/cellulose or
cellulose) was removed by centrifugation and 200 ml of the
supernatant was transferred to a fresh test tube. The KOH
supernatant (200 ml) was also neutralized and dialyzed to remove
the salts. All samples were dried, 25 mg myo-inositol was added as
an internal standard, and their glycosyl compositions were
determined by GLC of the alditol acetate derivatives of the
monosaccharides released by acid hydrolysis (see above).
Acknowledgements
This research was supported in part by funds from the US
Department of Energy grant DE-FG05-93ER20115 and DE-FG0993ER20097. We would like to thank Carl Bergmann for the pure
EPG, and Lene Anderson and Hans Peter Heldt Hansen from Novo
Nordisk, Denmark for the XEG-preparation and the PME.
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