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Special Focus Issue – Review
Xyloglucan and its Interactions with Other Components
of the Growing Cell Wall
Yong Bum Park and Daniel J. Cosgrove*
Department of Biology, 208 Mueller Laboratory, Pennsylvania State University, University Park, PA 16802, USA
*Corresponding author: E-mail, [email protected]; Fax, +1-814-865-9131.
(Received November 18, 2014; Accepted December 3, 2014)
The discovery of xyloglucan and its ability to bind tightly to
cellulose has dominated our thinking about primary cell wall
structure and its connection to the mechanism of cell enlargement for 40 years. Gene discovery has advanced our
understanding of the synthesis of xyloglucan in the past
decade, and at the same time new and unexpected results
indicate that xyloglucan’s role in wall structure and wall
extensibility is more subtle than commonly believed.
Genetic deletion of xyloglucan synthesis does not greatly
disable cell wall functions. Nuclear magnetic resonance studies indicate that pectins, rather than xyloglucans, make the
majority of contacts with cellulose surfaces. Xyloglucan
binding may be selective for specific (hydrophobic) surfaces
on the cellulose microfibril, whose structure is more complex than is commonly portrayed in cell wall cartoons.
Biomechanical assessments of endoglucanase actions challenge the concept of xyloglucan tethering. The mechanically
important xyloglucan is restricted to a minor component
that appears to be closely intertwined with cellulose at limited sites (‘biomechanical hotspots’) of direct microfibril
contact; these may be the selective sites of cell wall loosening by expansins. These discoveries indicate that wall extensibility is less a matter of bulk viscoelasticity of the matrix
polymers and more a matter of selective control of slippage
and separation of microfibrils at specific and limited sites in
the wall.
Keywords: Biomechanical hot spots Cellulose microfibrils Cell wall loosening Endoglucanase Expansin Pectins.
Abbreviations: AFM, atomic force microscopy; CEG, cellulose-specific endoglucanase; CSLC, cellulose synthase-like C;
CXEG, cellulose and xyloglucan hydrolyzing endoglucanase;
EM, electron microscopy; GPC, gel permeation chromatography; NMR, nuclear magnetic resonance; SEC-MALLS, size
exclusion chromatography coupled with multiangle laser
light scattering; ssNMR, solid-state NMR; XEG, xyloglucanspecific endoglucanase; XTH, xyloglucan endo-transglycosylase/hydrolase.
Introduction
Xyloglucans were initially identified as seed storage polysaccharides from nasturtium, tamarind and other seeds
(Kooiman 1957), and later as extracellular polysaccharides in
sycamore cell suspension cultures (Aspinall et al. 1969, Bauer
et al. 1973) and soon thereafter as components of primary cell
walls of many species (reviewed by Hayashi 1989). The concept
that xyloglucans play a central role in the control of wall extensibility originated with the pioneering molecular model of
the cell wall of sycamore cell suspension cultures by the
Albersheim group (Keegstra et al. 1973) and the discovery
that auxin-induced growth in pea epicotyls is accompanied
by increased xyloglucan metabolism (Labavitch and Ray
1974). Further evidence for xyloglucan binding to cellulose
(e.g. Hayashi and Maclachlan 1984), and for auxin-induced
changes in xyloglucan size (e.g. Nishitani and Masuda 1983,
Talbott and Ray 1992a), cemented xyloglucan as a focal element
in discussions of cell wall structure and growth for decades. The
view that ‘enzyme-catalyzed modification of xyloglucan crosslinks in the cellulose/xyloglucan network is required for the
growth and development of the primary cell wall’ (Pauly et al.
1999) became almost axiomatic in the cell wall field for more
than two decades. Hence the report of an Arabidopsis mutant
that lacked xyloglucan, yet exhibited only minor growth phenotype, came as a jolt to cell wall researchers (Cavalier et al. 2008).
This report, combined with a number of other results at odds
with current models, has led us and some other researchers to
rethink the structure of primary cell walls and the role of xyloglucan in cell wall extensibility.
In the 40 years since xyloglucan came to be viewed as an
indispensable component controlling wall extensibility, the concepts of how xyloglucan interacts with cellulose and other wall
components have evolved considerably. The original hypothesis
of a macromolecular matrix made of covalently linked domains
of xyloglucan, pectins and structural proteins (Keegstra et al.
1973) was replaced by the simpler tethered network model
(Hayashi 1989, Carpita and Gibeaut 1993, Nishitani 1998,
Cosgrove 2001) which highlighted direct coating and tethering
of cellulose by xyloglucan as the key structural determinant of
wall extensibility. Despite widespread acceptance of this model,
numerous points in the model have not been critically tested and
it is probably too tight-knit a structure to be compatible with the
high mechanical extensibility and flexibility of many primary cell
walls (Hepworth and Bruce 2004). Some alternative models lacking direct cellulose–cellulose linkages have been suggested; for
instance, Talbott and Ray (1992b) proposed a ‘multicoat’ structure in which cellulose was coated with tightly bound xyloglucan,
which was coated with more loosely held arabinans/galactans,
Plant Cell Physiol. 56(2): 180–194 (2015) doi:10.1093/pcp/pcu204, Advance Access publication on 21 January 2015,
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Plant Cell Physiol. 56(2): 180–194 (2015) doi:10.1093/pcp/pcu204
which were intermingled with a gelled interface of acidic polysaccharides and structural protein; Thompson (2005) suggested
that wall extensibility is controlled by spatial constraints in the
movement of microfibrils rather than by direct interfibril linkages. These authors noted important shortcomings in the popular model, but their alternative concepts have garnered little
support or follow-up.
Recent results based on solid-state nuclear magnetic resonance (ssNMR) and biomechanical approaches have raised fresh
doubts about the correctness of the tethered network model
and, more generally, about the significance of xyloglucan’s role
in wall structure and mechanics. Concomitantly, pectin has
seen a resurgence as an important element in discussions of
cell wall mechanics (Zykwinska et al. 2008, Palin and Geitmann
2012, Braybrook and Peaucelle 2013). Thus, this seems to be a
good juncture to review recent developments in the field of
xyloglucan and to assess their implications for cell wall structure and extensibility as well as to point out areas in need of
more in-depth inquiry.
This review considers xyloglucan function in the context of
common models (or, more correctly, depictions) of growing cell
walls: cellulose microfibrils are represented as well-spaced and
non-contacting rods; xyloglucan covers most cellulose surfaces,
preventing direct contact between microfibrils and simultaneously acting as the major (or sole) load-bearing tether between
cellulose microfibrils; wall strength stems from xyloglucan
tethers between adjacent microfibrils and wall loosening depends on breaking or shifting the xyloglucan tethers. These
concepts of xyloglucan function emerged from a large body
of work, cogently summarized in various reviews (Fry 1989,
Hayashi 1989, Carpita and Gibeaut 1993, Cosgrove 2001) and
extended by later results, including transgenic expression of
xyloglucanase and cellulase in plant tissues (Park et al. 2003,
Hayashi and Kaida 2011). The tethered network is the most
common depiction in overviews of plant cell walls (e.g.
Somerville et al. 2004, Cosgrove 2005, Burton et al. 2010,
Scheller and Ulvskov 2010), but many of its features and implications are untested. In this article, we emphasize some of the
more physical aspects of xyloglucan–cellulose interaction that
are not often discussed in the biological literature, we review
recent results that appear to run counter to these wellentrenched concepts of primary walls, and we suggest possible
ways to resolve some apparent contradictions.
We begin with a brief update on xyloglucan structure and
synthesis, then summarize studies on the physical binding of
xyloglucan to cellulose, followed by recent physical, genetic and
biomechanical tests of the tethered network model. We do not
discuss the potential role of xyloglucans in secondary cell walls
(but see Mellerowicz et al. 2008, Baba et al. 2009, Hayashi and
Kaida 2011, Nishikubo et al. 2011).
Structure, Size, Diversity and Conformation of
Xyloglucans
Xyloglucans comprise a heterogeneous collection of polysaccharides of variable length and side chain pattern. They are
widespread if not universal in the primary cell walls of land
plants, but their amounts vary considerably. In many dicots,
xyloglucans constitute the major hemicellulose of growing cell
walls, comprising approximately 20% of the dry mass of primary
cell walls (reviewed in Fry 1989, Hayashi 1989, Schultink et al.
2014), but xyloglucan content may be as low as 2% (Thimm
et al. 2002). Grasses—but not monocots in general—have a
reduced xyloglucan content; values of approximately 5% of primary walls are typical in grasses, but values as high as 10% occur
(Carpita 1996, Gibeaut et al. 2005). Xyloglucans consist of a b(1,4)-D-glucan backbone that is quasi-regularly substituted with
a-D-xylosyl residues linked to glucose through the O-6 position
(Fig. 1A, B). In many species, the backbone has a regular pattern of three substituted glucose units followed by an unsubstituted glucose residue. Substitution is less frequent in some
taxonomic groups. For instance, only about 40% of the glucose
residues are substituted in tomato and other species in the
Solanaceae (Ring and Selvendran 1981, York et al. 1996) while
xyloglucans from grass cell walls have 30–40% substitution
(Hsieh and Harris 2009).
The xylosyl residue may be b-(1,2) linked with a D-galactosyl
residue which may be additionally linked a-(1,2) with an
L-fucosyl residue; other side chain patterns have also been documented (Schultink et al. 2014, Tuomivaara et al. 2015). A concise nomenclature is used as shorthand for the side chain
structure (Fig. 1D). Diagnostically useful endoglucanases cut
the xyloglucan backbone selectively at the unsubstituted position, producing a mixture of oligosaccharides commonly made
of four glucose residues in the backbone with three xylose and a
smaller number of galactose and fucose substituents. These
oligosaccharides are readily identified by high-performance
anion exchange chromatography with pulsed amperometric
detection or matrix-assisted laser-desorption/ionization time
of flight mass spectrometry, which have been adapted for sensitive and rapid xyloglucan analysis (Gunl et al. 2011,
Tuomivaara et al. 2015).
In many dicots, the most common oligosaccharides produced by endoglucanase digestion of xyloglucan are the Glc4
oligosaccharides designated XXXG, XXFG, XXLG and XLFG, but
numerous species-dependent and organ-dependent variations
in the side chain pattern have been documented (Schultink
et al. 2014, Tuomivaara et al. 2015). For instance, galactose is
frequently replaced by arabinose in xyloglucans of tomato and
other species in the Solanaceae (Ring and Selvendran 1981,
Hoffman et al. 2005), whereas galacturonic acid partially replaces galactose in an unusual acidic xyloglucan recently identified in Arabidopsis root hairs (Pena et al. 2012). The
xyloglucan side chains in grasses are generally not extended
beyond xylose; upon endoglucanase digestion, the most
common oligosaccharides are XXGG and XXGGG. Thus, the
defining structure for xyloglucan is a glucan backbone with
O-6-linked xylose side chains, but the extent of further substitution and sugar identity varies considerably. As discussed
below, side chains may affect interactions with cellulose and
possibly other components of the cell wall, but whether such
interactions play a significant functional role in the biological
variation of xyloglucan is not entirely clear at this point.
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Fig. 1 Xyloglucan structure. (A) Structure of the XLLG oligosaccharide, showing the b-1,4-D-glucan backbone (gray) with side chains made of
xylose (green) and galactose (blue). (B) Pattern of linkages for sugars in the XLFG oligosaccharide with the associated glycosyl transferases
indicated in upper case letters. (C) Xyloglucans may have short regions in a flat-ribbon conformation, suitable for binding to the hydrophobic
surface of cellulose microfibrils or may assume a twisted conformation. (D) List of naturally occurring xyloglucan side chains and their one-letter
designations. Image credits: B is from Zabotina (2012), reprinted under Creative Commons Attribution Non Commercial License; C is reprinted
from O’Neill and York (2003), copyright Wiley-Blackwell, used with permission. D is adapted from Tuomivaara et al. (2014), copyright by Elsevier
and used with permission.
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Another notable feature of XyG is O-acetylation, which is
often overlooked because the ester linkage is broken by the
strong alkaline conditions commonly used to extract xyloglucan from cell walls. In Arabidopsis and many other dicots,
O-acetylation of XyG is found principally on galactosyl residues
at the O-3 or O-2 positions (Kiefer et al. 1989). This contrasts
with xyloglucans in grasses (Poaceae) and the Solanaceae where
the glucan backbone is acetylated at O-6, in effect replacing the
xylose side chain with an acetyl group (Gibeaut et al. 2005, Jia
et al. 2005). Acetylation was not detected in xyloglucans from
apple pulp or tamarind seeds (Sims et al. 1998), indicating biological variation in this structural trait. Acetylation of galactose
is not essential in Arabidopsis, as at least one ecotype lacks it,
but it may contribute to pathogen resistance (Manabe et al.
2011, Gille and Pauly 2012). Acetylation of the substituted glucose residues in xyloglucans of the Poaceae and the Solanaceae
improves xyloglucan solubility and prevents self-association
and aggregation in these xyloglucan variants that have relatively
low substitution (Sims et al. 1998). One might surmise that
grass xyloglucans bind more tightly to cellulose than do the
more highly substituted xyloglucans of other plant groups
(see below), but this issue has evidently not been examined
closely.
Chain length
Published size estimates of xyloglucan from primary cell walls
are based mostly on gel permeation chromatography (GPC; also
known as gel filtration chromatography or size exclusion chromatography). Peak values for relative molecular mass (Mr) vary
enormously, ranging from 9 kDa to >900 kDa, with most
estimates in the 100–300 kDa range; moreover, xyloglucan is
reported to undergo rapid shifts in Mr under various conditions
(Nishitani and Masuda 1983, Talbott and Ray 1992a, Talbott
and Pickard 1994, Thompson and Fry 1997). Storage xyloglucans, i.e. from tamarind, nasturtium and other seeds, are exceptionally large, reported to exceed 1,000 kDa (Lima et al. 2004).
The smallest values (9 kDa) (Talbott and Ray 1992a) correspond
to a backbone length of 28 glucose residues (14 nm) whereas
900 kDa corresponds to 2,800 glucose residues (1,400 nm). This
100-fold range in peak values is truly remarkable for a wall
component thought to play a central structural role in cell
wall architecture and mechanics.
Chain length estimates based on GPC must be regarded with
caution because they are sensitive to the conformation, hydrodynamic radius and aggregation of the polymer, as well as to
interactions with the column matrix and other potential influences. Moreover, iodine staining, which is used to quantify
xyloglucan, may bias the results toward the large size because
it is insensitive to xyloglucans <10–20 kDa (Hayashi 1989,
O’Neill and York 2003). GPC columns are typically calibrated
with dextrans [a-1,6-D-glucan backbones with 5% a-(1,3)glucose substitution] which have a different conformational
behavior from xyloglucans, potentially introducing large overestimates in Mr. This issue is generally well recognized
(Talbott and Ray 1992b, Gaborieau and Castignolles 2011,
Kohnke et al. 2011) and such Mr data are best interpreted
as relative rather than absolute size. Size exclusion
chromatography coupled with multiangle laser light scattering
(SEC-MALLS) can provide better estimates of polymer size
(Harding 2005), but this method has rarely been employed
for xyloglucan from primary cell walls.
An appreciation of the problems of chain length estimates
based on GPC alone can be gained from a study of wheat
arabinoxylan that was partially digested with arabinosidase to
remove some of the side chains but to leave the backbone
intact. The hydrodynamic radius of the polymer shrank from
24 nm to 9 nm as the percentage of unsubstituted backbone
residues increased from 63% to 84% (Kohnke et al. 2011). Using
dextrans for calibration (Armstrong et al. 2004), one would
estimate a 10-fold reduction in polymer length from these
values, but in this instance the difference was primarily due
to conformation because arabinose removal reduced the molecular mass by only 11%. The polymer with less substitution
assumed a more compact conformation; self-aggregation of the
backbone may have contributed to the smaller size. In the case
of xyloglucans, conformation may vary with the degree of substitution, the length of side chains and the acetylation pattern;
xyloglucans may also be modified during extraction and may
undergo changes in aggregation and association with other
substances that affect their elution from a GPC column. Most
GPC-based estimates of xyloglucan Mr probably lead to overestimates of xyloglucan chain length.
A very different approach to estimating chain length makes
use of electron microscopy (EM) or atomic force microscopy
(AFM) to measure the length of extracted polymers dispersed
on a smooth mica surface. These methods do not reveal the
chemical identity of the polymers, which must be verified by
other means. An EM analysis of presumptive xyloglucans (linear
polymers from a 1 M KOH extract of onion scale parenchyma
walls) showed chain lengths ranging from 20 to 300 nm, with a
number-averaged length of approximately 150 nm (McCann
et al. 1992). Assuming these were indeed single xyloglucan molecules, not aggregates or concatenated chains, they would correspond to an average xyloglucan size of approximately 90 kDa
(number-average estimate; the mass-average estimate would be
larger). This study also noted a stepwise distribution of xyloglucan chain lengths, with steps of approximately 30 nm. The basis
for the steps is uncertain, but it suggests a mechanism of blockwise assembly or restructuring of xyloglucan in muro, i.e. by
xyloglucan endo-transglucosylase (Nishitani and Tominaga
1992, Thompson and Fry 1997, Rose et al. 2002). Microscopic
measurements of polymer chain lengths are labor intensive and
have their own set of technical problems, but they provide
detailed information not provided by other methods. This appears to be the only study of xyloglucan size distribution by this
method up to now. AFM has been used to image tamarind
xyloglucan dried on mica, but without detailed length analysis
(Morris et al. 2004); it has also been used to quantify the
distribution of pectin chain lengths (Round et al. 2010).
Different biological sources undoubtedly contribute to the
wide range of xyloglucan Mr values, but even with the same
material (third internode of etiolated pea epicotyl) the major
xyloglucan peak was estimated in one study to be approximately 300 kDa (Hayashi and Maclachlan 1984) and 30 kDa in
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another (Talbott and Ray 1992b). The difference was attributed
to rapid and reversible changes in xyloglucan size induced by a
variety of biological treatments (Talbott and Ray 1992a), but
the molecular basis for such reversible changes has not been
established.
After reviewing the xyloglucan literature, we must admit to
being impressed by the striking variation in peak Mr values for
xyloglucans in growing cell walls. Can they really vary dynamically by >10-fold? How do such changes influence cell wall
extensibility? A common view is that smaller xyloglucans
mean greater wall extensibility, e.g. as expressed by Fry
(1989), but a well-controlled study of this point has not been
published. The observations of Talbott and Ray (1992a) challenge a simple correlation between Mr and extensibility, as do
recent results with substrate-specific endoglucanases, to be discussed below.
Another issue concerns xyloglucan conformation in the wall.
At short length scale (10 glucose residues) the xyloglucan
backbone assumes an extended, relatively rigid conformation
(Fig. 1C) whose flexibility depends on the side chains and their
interaction with the backbone (Gidley et al. 1991, Levy et al.
1997), whereas at longer scale the polymer in solution forms a
‘semiflexible swollen coil’ (Muller et al. 2011). Small-angle neutron scattering measurements indicate that xyloglucan in solution has a persistence length of 8 nm (four Glc4 units or 16
glucose residues). Other techniques give similar or somewhat
smaller values (Patel et al. 2008). Persistence length is a measure
of polymer rigidity; roughly, it is the distance over which a chain
will bend significantly (or lose orientational correlation) by
thermal fluctuations. For comparison, the DNA double helix
has a persistence length of 50 nm, i.e. it is a much more rigid
molecule, whereas dextran has a much smaller persistence
length, estimated as 0.4 nm (Rief et al. 1998). Well-solvated,
self-avoiding polymers whose length is much longer than the
persistence length will occupy a time-averaged spheroidal
shape with a diameter much less than the chain length. In
the case of tamarind xyloglucan, the hydrodynamic radius
was measured as 51 nm and its weight-average molecular
mass was determined by SEC-MALLS to be 470 kDa, corresponding to approximately 750 nm extended chain length
(Muller et al. 2011). These results show that xyloglucan in solution assumes a coiled shape, characteristic of most polymers,
but stiffer than highly flexible chains such as dextran. The shape
of xyloglucans in solution thus differs from the extended conformation represented in many depictions or ‘cartoons’ of primary cell walls. It may be argued that xyloglucan–cellulose
interactions would force xyloglucan into an extended conformation, but NMR results, discussed below, suggest that only a
small fraction of xyloglucan in the wall has such a conformation.
A final structural issue is the extent to which xyloglucan is
covalently cross-linked to other components of the cell wall, as
well as the (unknown) nature of the putative covalent linkage.
There are reports that up to 30% or more of the xyloglucan in
walls from cell suspension cultures is covalently linked to
pectin, evidently cross-linked during synthesis within the
Golgi apparatus (Thompson and Fry 2000, Popper and Fry
2008). In contrast, numerous studies of cell walls from growing
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tissues (not cell cultures) show that little or none of the xyloglucan is covalently linked to pectins (see Talbott and Ray
1992b, and references therein). A simple method dubbed ‘epitope detection chromatography’ was recently combined with a
simple sample preparation procedure to characterize polysaccharide complexes from Arabidopsis cell walls (Cornuault et al.
2014). The method passes solubilized components through an
anion exchange column and uses antibodies to detect specific
epitopes in the eluted fractions. The results indicated that the
vast majority of xyloglucan was not linked to pectin, judged by
lack of retention on the anion exchange column. In extracts
from shoots, the xyloglucan–pectin component amounted to
only a trace, while root extracts showed a notable but still
minor peak, approximately 5% of the total, that is potentially
a xyloglucan–pectin hybrid. These results are consistent with
most biochemical dissections of primary cells walls. They contrast profoundly with a study of xyloglucans from Arabidopsis
cell cultures where 50% of the xyloglucans appeared to be
synthesized on a pectin primer, forming a chimeric polysaccharide that was stable in the cell wall for several days (Popper and
Fry 2008).
How can such divergent results be understood? Perhaps the
ability to link xyloglucan and pectin intracellularly is adaptively
expressed under special circumstances and may be particularly
well developed in cell suspension cultures. Cell cultures differ
from cells in the plant body and are capable of remarkable
adaptations in wall structure (Shedletzky et al. 1990).
Xyloglucan–pectin complexes might be an intermediate stage
in the synthesis of specific wall components (Tan et al. 2013),
followed by polymer rearrangements in the cell wall mediated
by transglycosylases or other lytic enzymes. Cross-linking of
xyloglucan with pectin also offers a potential mechanism for
rigidifying cell walls as they cease growth and become insensitive to wall-loosening agents such as a-expansin (Cosgrove
1996, Zhao et al. 2008). We conclude that chimeric xyloglucan–pectin molecules are quantitatively minor components
of most growing cell walls. This does not necessarily mean
they are insignificant for wall integrity or wall extensibility,
but this point has not been critically tested to date. A major
step toward this goal would be to identify the enzyme that
forms the putative linkage between the two polymers, followed
by biomechanical analysis of walls from mutants with defects in
the corresponding gene.
Biosynthesis
Identification of the genes underlying xyloglucan biosynthesis
has progressed rapidly in recent years (Zabotina 2012, Zabotina
et al. 2012 Pauly et al. 2013) and is only briefly recapped here
(Fig. 1B). The b-(1,4)-D-glucan backbone of XyG is synthesized
by Golgi-localized glycan synthases, encoded by CELLULOSE
SYNTHASE-LIKE C (CSLC) genes (Cocuron et al. 2007). Side
chains are added in the Golgi by multiple glycosyltransferases
including a-(1,6)-xylosyltransferases (XXT1, XXT2 and XXT5),
b-(1,2)-galactosyltransferases and a-(1,2)-fucosyltransferase
(Perrin et al. 1999, Faik et al. 2002, Vanzin et al. 2002, Madson
Plant Cell Physiol. 56(2): 180–194 (2015) doi:10.1093/pcp/pcu204
et al. 2003, Pena et al. 2004, Lerouxel et al. 2006, Zabotina et al.
2008, Zabotina et al. 2012). Moreover side chain substitution is
finely tuned by different glycosyltransferases that add the same
glycosyl moiety to different positions (Schultink et al. 2014).
The transfer of galacturonic acid to xylose in Arabidopsis root
hairs appears to be catalyzed by a family-47 glycosyltransferase
(Pena et al. 2012). CSLC4 and some of the xylosyltransferases
form a multiprotein synthesis complex in the Golgi (Chou et al.
2012; see also the review in Zabotina 2012). To create novel
xyloglucans, Schultink et al. (2013) expressed foreign arabinosyltransferases in a mur3/xlt2 Arabidopsis mutant lacking xyloglucan galactosylation. This resulted in a novel xyloglucan that
was intermediate in structure between that of Arabidopsis and
Solanaceous species. The mur3/xlt2 mutants, which produced
xyloglucan made only of XXXG subunits, were shorter than the
wild type and had reduced wall extensibility. This phenotype
may stem from dysfunctional xyloglucans within the wall or
from detrimental effects on Golgi function (Tamura et al.
2005). The transgenic plants produced xyloglucan containing
XXSG subunits (S indicates a xylose-arabinose side chain). This
change restored growth and wall extensibility to wild-type
levels. The results suggest that the length of the xyloglucan
side chain may be more important than the specific type of
sugars used. Thus the era of ‘designer xyloglucans’ is upon us,
and this platform may prove useful in the future for creating
additional novel xyloglucan variants to explore more detailed
structure–function relationships of xyloglucan.
Complementary to these biosynthetic enzymes, plants also
have a suite of glycosidases that can trim xyloglucan side chains
after deposition to the cell wall (Pauly et al. 2001, Sampedro
et al. 2010). Such enzymes may be involved in turnover or
recycling of sugars from non-structural xyloglucans in the cell
wall.
As mentioned above, galactose residues in xyloglucan are
acetylated in Arabidopsis (Gille and Pauly 2012). Forward genetic approaches identified a pair of related multidomain transmembrane proteins named AXY4 and AXYL as essential for
galactose acetylation (Gille et al. 2011). These proteins are
part of a flexible O-acetylation mechanism that transfers
acetyl groups to a number of polysaccharides and that is conserved across kingdoms (Gille and Pauly 2012). The targeting of
acetylation, e.g. to the glucan backbone in xyloglucans of
grasses or solanaceaous species, or to the galactose side chain
in Arabidopsis and other species, or to other targets, is apparently well regulated but not understood in detail.
One notable disparity in the literature is the remarkably
small size of xyloglucans isolated from the Golgi (9 kDa)
(Talbott and Ray 1992a, Thompson and Fry 1997) compared
with the much larger size of xyloglucans extracted from cell
walls (Mr in the range of 100–300 kDa or larger). One possible
explanation for this disparity is that xyloglucan is synthesized in
the Golgi as smaller units that are secreted to the wall and
subsequently linked together to make longer chains. Such ligations are possible through the action of members of the xyloglucan endo-transglucosylase/hydrolase (XTH) family
(Thompson et al. 1997, Thompson and Fry 2001, Rose et al.
2002), potentially generating the stepwise distribution of
xyloglucan lengths observed by McCann et al. (1992). The
same enzyme might also underlie other dynamic shifts in xyloglucan size (Talbott and Ray 1992a, Talbott and Pickard 1994),
but this has not been established. If mature xyloglucans are
indeed enzymatically assembled in the cell wall from small segments, the process could have a major impact on the topology
of xyloglucan within the wall, probably resulting in complex
entanglements; this could explain the need for a high NaOH
concentration to extract xyloglucan from cell walls, whereas
xyloglucan bound to cellulose in vitro is released at much
lower concentrations.
Interactions of Xyloglucan with Cellulose In
Vitro and In Silico
Early studies established that xyloglucan bound tightly but noncovalently to cellulose (Aspinall et al. 1969, Bauer et al. 1973).
This was confirmed and extended by later studies, eventually
leading to the ‘tethered network’ concept that xyloglucan extensively coats cellulose surfaces and tethers microfibrils together (Hayashi 1989, Pauly et al. 1999). Xyloglucan binding
in vitro to cellulose has been explored with several physical
and biological techniques. Many such studies have used commercially available tamarind seed xyloglucan, which differs in
size, acetylation and side chain composition from xyloglucans
typical of primary cell walls. Moreover, the forms of cellulose
used for in vitro studies often differ from native cellulose from
primary cell walls. It has become evident that these structural
differences could impact the applicability of the results for
understanding the state of xyloglucans in native cell walls. For
instance one study found that the relative binding capacity for
fucosylated and de-fucosylated xyloglucans differed according
to cellulose source (Chambat et al. 2005). The two forms of
xyloglucan bound equally to cellulose prepared from primary
cell walls, but binding differed 2-fold with cellulose from secondary cell walls of flax. The lesson is clear: the biological source
of cellulose and its processing can substantially influence xyloglucan binding.
Native cellulose from primary cell walls takes the form of
3 nm wide microfibrils, consisting of cellulose crystalline domains and less ordered regions (Thomas et al. 2013, Cosgrove
2014). The degree of crystallinity is low, with estimates as low as
20–40% (Chambat et al. 2005). This contrasts with celluloses
from secondary cell walls or bacterial sources which have much
higher crystallinity and may aggregate into larger ordered structures. The crystalline regions have surfaces with distinctive
physical properties: surfaces with exposed sugar rings form a
dominantly hydrophobic planar surface populated by axial CH
groups, whereas the other surfaces are populated by equatorial
OH groups extending from the sides of the stacked glucan
chains, and consequently are hydrophilic (Fig. 2A). The two
surfaces probably interact in different ways with water, matrix
polymers, enzymes and small molecules. As illustrated in Fig. 2,
the ratio of hydrophilic to hydrophobic surface depends on the
shape of the microfibril cross-section. This is an important,
but poorly characterized, aspect of microfibril structure
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Y. B. Park and D. J. Cosgrove | Xyloglucan interactions in growing cell walls
Fig. 2 Cross-sectional shape and width of cellulose microfibrils affect
the relative surface area with hydrophobic (blue) and hydrophilic
(yellow) character. Left: a 24 chain microfibril with a truncated diamond shape. Right: an 18 chain microfibril with a rectangular shape. A
xyloglucan chain is shown bonded non-covalently in an extended
conformation on the upper hydrophobic surface of each microfibril.
(Newman et al. 2013). Presumably the same kinds of surfaces
are found in non-crystalline regions of cellulose, but there
seems to be little evidence regarding this point. Two crystalline
allomorphs (cellulose Ia and Ib) are synthesized by plants, algae
and bacteria. They differ slightly in the alignment of adjacent
glucan chains in the crystal. Land plants produce predominantly the cellulose Ib form, while algae and bacteria produce
mostly cellulose Ia (Atalla and Vanderhart 1984, Lee et al. 2014),
but the functional significance of this difference is unclear.
Although celluloses from different sources are chemically
identical, their physical forms may differ, potentially influencing
xyloglucan binding (Chambat et al. 2005). The degree of crystallinity and the specific surface area are key factors, but other
aspects may also be involved. Regenerated cellulose surfaces
and amorphous cellulose particles lack the repeating organized
structure of the cellulose microfibril. So-called amorphous cellulose, produced in vitro by swelling or dissolving crystalline
cellulose, probably differs in surface structure from the noncrystalline portion of cellulose found in primary cell walls. A
commonly used form of ‘microcrystalline cellulose’ branded as
Avicel contains crystalline regions of Ib cellulose, but its physical form is an irregular, porous particle of acid-washed, pulverized woody cell wall aggregates (Supplementary Fig. S1).
Smaller xyloglucan polymers can penetrate deeper into the
Avicel particle than can larger polymers, which are restricted
to the larger pores. Because the effective surface area for binding varies with xyloglucan size, binding kinetics are complicated
and partly limited by size-dependent xyloglucan diffusion into
the pores of the particles. These geometrical issues probably
account for the markedly higher binding of Avicel for smaller
xyloglucan polymers reported by Lima et al. (2004). In studies
that used less porous cellulose, the binding capacity showed
little dependence on xyloglucan size or increased with the size
of the xyloglucan chain (Hayashi et al. 1994, Lopez et al. 2010),
indicative of a surface topology with loops and coils.
To obtain forms of cellulose more closely resembling native
surfaces, cellulose has been isolated from primary walls (Hayashi
et al. 1987), but treatment with strong alkali to remove hemicelluloses would have caused swelling of cellulose and
186
converted it to a different crystalline isomorph (cellulose II),
in which the glucan chains are said to run antiparallel rather
than parallel as in the native form (cellulose I). Hydrolysis with
HCl has also been used (Chambat et al. 2005). This treatment
may avoid the swelling problem, but it hydrolyzes some noncrystalline regions of the microfibrils, producing more tractable
cellulose nanofibrils or nanocrystals which may partially resemble primary cell wall cellulose, but probably differ in some structural aspects. In other studies, nanocrystalline cellulose has
been made by digestion with sulfuric acid, which is problematic
because it esterifies the surface cellulose chains with sulfate
groups, leaving a large negative charge on the surface
(Gu et al. 2013). The sulfate esters may be removed by subsequent treatment with NaOH. These practical complexities of
cellulose structure do not negate the value of in vitro binding
studies but they do need to be kept in mind in evaluating the
applicability of results for native wall structure.
When applied in dilute aqueous solutions, xyloglucans bind
to cellulose as a monolayer, meaning that xyloglucan does not
stack on itself; the binding appears to be irreversible and the
binding capacity—measured as the xyloglucan : cellulose
ratio—increases as the fibril size is reduced, as would be expected for the trend of surface area : mass ratio (Hayashi et al.
1987, Chambat et al. 2005). Much attention has been given to
the significance of terminal fucose residues on trisaccharide (F)
side chains since an early computational study suggested that F
side chains promote a xyloglucan conformation favorable for
cellulose binding (Levy et al. 1991). Experimental work showed
that fucosylated xyloglucan bound more rapidly to Avicel than
did non-fucosylated xyloglucan (Hayashi et al. 1994, Levy et al.
1997), but a reverse trend was observed with other forms of
cellulose (Chambat et al. 2005). These xyloglucans were isolated
by extraction with alkali, and so would be de-acetylated.
Xyloglucans bound to cellulose in vitro are readily solubilized
with 0.25 M NaOH, which is much lower than the 4 M concentration required for extraction from native cell walls. This difference in extractability presumably stems from the more
intricate interactions/entanglements of xyloglucans with cellulose and pectins in native cell walls, arising from the wall assembly process. Such interactions have been partially mimicked
in multilayered assemblies of cellulose nanocrystals and
xyloglucan (Cerclier et al. 2010) and in cellulosic composites
produced by cellulose-synthesizing bacteria grown with
xyloglucan in the culture medium (Whitney et al. 2006).
By use of adsorption isotherms and isothermal titration calorimetry, a recent study explored the influence of chain length,
side chain pattern and degree of acetylation on xyloglucan
binding to nanocrystalline cellulose (Lopez et al. 2010).
Celluloses were prepared from bacterial cellulose or filter
paper by acid hydrolysis, so aspects of their structure may
differ from cellulose in primary cell walls. Two stages of xyloglucan adsorption to cellulose were identified: an exothermic
process attributed to hydrogen bonding at low xyloglucan concentration, and an endothermic process attributed to hydrophobic interactions and solvent reorganization at high
xyloglucan oligosaccharide concentration. This study found
that binding capacity increased with the size of the xyloglucan,
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that a minimum of 12 backbone glucosyl residues were needed
for binding to crystalline cellulose surfaces, and that xyloglucan
with fucose-terminated side chains bound at somewhat higher
levels than did xyloglucan lacking fucose. These results are consistent with previous work (Hayashi et al. 1994). The minimum
chain length for binding is similar to the persistence length of
xyloglucan, suggesting that binding requires bonding between a
relatively stiff xyloglucan segment and a flat cellulose surface.
The enhanced binding of fucosylated xyloglucan was attributed
to the trisaccharide character of the fucose-containing side
chain—potentially providing more OH groups for hydrogen
bonding to cellulose—rather than any particular influence of
fucose on xyloglucan conformation. This inference was based
on a computational analysis of xyloglucan–cellulose interactions (Hanus and Mazeau 2006) rather than experimental
results. Acetylation did not influence xyloglucan binding to
nanocrystalline cellulose.
Lopez et al. also noted that xyloglucan binding did not involve a large change in enthalpy (heat), confirming an earlier
study (Lima et al. 2004). The low enthalpic character of binding
suggests that binding has a large entropy component, most
probably by freeing water molecules constrained to an organized layer of water at cellulose surfaces. The entropy contributions for xyloglucan–cellulose binding require further study,
especially in view of the crowded environment of the cell wall
where molecular crowding might influence polymer interactions (Benton et al. 2012). It is notable in this context that
amorphous cellulose is reported to have more hydrophilic and
fewer hydrophobic surfaces exposed to aqueous solvent than
does crystalline cellulose (Mori et al. 2012). This observation
may be relevant to the mode of xyloglucan binding in primary
cell walls where the majority of cellulose is microfibrillar but
with low crystallinity.
Xyloglucan binding to the distinctive surfaces of cellulose is
difficult to distinguish experimentally, but insights may be
gained from computational approaches. Early computational
work simulated xyloglucan binding to cellulose without explicitly considering the contribution of water (Levy et al. 1991,
Finkenstadt et al. 1995, Levy et al. 1997, Hanus and Mazeau
2006). In contrast, more recent studies included water and
used different quantitative approaches based on more
advanced molecular dynamics software (Zhang et al. 2011,
Zhao et al. 2014), reaching somewhat different conclusions
from the earlier work and showing that the influence of side
chains on binding is more complicated than anticipated in
earlier work. Inclusion of water substantially reduced the
interaction energy calculated in the binding simulations. With
xyloglucan oligosaccharides (glucan chain length = 9) bound to
a hydrophilic surface of cellulose, galactosyl and fucosyl
substitutions had little effect on binding strength, but xyloglucan conformation was affected (Zhang et al. 2011). When interaction energies were compared for xyloglucan bound to the
hydrophilic and hydrophobic surfaces of cellulose in water,
binding was seen to be much more favorable for the hydrophobic surfaces for most side chain configurations tested (Zhao
et al. 2014). Such high-energy binding entailed removal of structured water from the cellulose surface and from half of the
xyloglucan surface. Presumably such structured water pays an
entropic penalty which would be redeemed upon its release by
xyloglucan binding to the cellulose surface. Xyloglucan was
much more rigid when bound to the hydrophobic surface
than when bound to the hydrophilic surface, where binding
was more disorderly and of lower energy. This computational
study suggests a geometrical specificity of binding to cellulose
surfaces that could be significant for the assembly of cell walls
in vivo. In a similar vein, surface dehydration and hydrophobic
interactions are considered important aspects of the formation
and insolubility of cellulose microfibrils (Glasser et al. 2012,
Medronho et al. 2012) and for binding of carbohydrate-binding
modules to the hydrophobic surface of cellulose microfibrils
(McLean et al. 2002, Georgelis et al. 2012).
Another computational study based on coarse-grain modeling suggested that xyloglucan binding to cellulose would create
tension in the polymer, i.e. tethered xyloglucans were seen to be
self-tensioning (Morris et al. 2004). O’Neill and York (2003)
proposed that the straightening and untwisting of xyloglucans
tethered in two places would drive the polymer into a duplexed, coiled conformation. Self-tensioning and duplex formation has not been experimentally verified, but perhaps they
contribute to force generation in tension wood (Baba et al.
2009). If such a self-tensioning process functioned in primary
cell walls, tethers would adaptively unzip and zip (lengthen and
shorten) to maintain a constant tension—an interesting concept that presents problems if xyloglucan function is to resist
wall stresses originating from cell turgor pressure. Would typical
wall stresses simply pull xyloglucan tethers off the cellulose
surface, as occurs during single molecule AFM spectroscopy
(Morris et al. 2004)? This possibility has not yet been assessed.
Other physical studies indicate that xyloglucan has two apparently contradictory effects on cellulose composites in vitro:
it reduces friction between cellulose surfaces while also increasing adhesion between adjacent fibril surfaces (Stiernstedt et al.
2006, Cerclier et al. 2010). These opposing effects depend on the
duration of the interaction and the distances between cellulose
surfaces. When xyloglucan was incorporated into bacterial
composites, the material became very soft and extensible
(Whitney et al. 2006); this effect was partly due to reduced
friction between cellulose fibrils but mostly it was a consequence of large xyloglucan-organized domains of cellulose fibrils that could readily slip past one another. Slippage of large
domains encounters less frictional resistance than nanoscale
slippage of many randomly entangled fibers. These methods
can reveal potential interactions that might occur in plant
cell walls, but the differences in materials, structure and environment from bona fide cell walls may limit extrapolation of the
conclusions to the biological realm where more organized
assembly processes may dominate wall structure.
Xyloglucan–Cellulose Interactions in the
Primary Cell Wall
We turn now to a more biological context. As described above,
a long-standing view holds that xyloglucan binds tightly to
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Y. B. Park and D. J. Cosgrove | Xyloglucan interactions in growing cell walls
cellulose within the primary cell wall, coating most available
cellulose surfaces, intercalating into microfibrils during their
synthesis and tethering adjacent microfibrils to form the
major load-bearing network in the growing cell wall. These
concepts, illustrated in many depictions of the cell wall, are
based upon several facts and assumptions (Hayashi 1989,
Mccann et al. 1992): (i) that xyloglucan binds tightly to cellulose
surfaces; (ii) that the amount of xyloglucan in primary walls is
sufficient to coat most or all accessible cellulose surfaces (this
statement assumes xyloglucan has a fully extended conformation throughout its length); (iii) that a large portion of xyloglucan in the cell wall is recalcitrant to extraction except under
conditions that swell cellulose, e.g. 4 M NaOH; (iv) that unextracted cell walls lack available sites for binding exogenous
xyloglucan; (v) that direct cellulose–cellulose contacts do not
occur in primary cell walls (a common assumption, but see
discussion below); and (vi) that xyloglucan forms interfibril
tethers with sufficient mechanical strength to withstand the
tensile forces generated by turgor pressure. We first consider
the concept that xyloglucan coats cellulose microfibrils, then
assess the concepts that it intercalates within microfibrils and
tethers microfibrils together.
Coating microfibrils
Contrary to the prediction of the conventional model, 13CssNMR analyses of complex cell walls showed that surprisingly
little of the cellulose microfibril surface makes contact with
xyloglucan (Bootten et al. 2004, Dick-Perez et al. 2011). In the
study by Bootten et al., proton spin relaxation was combined
with spectrum editing to identify xyloglucans of two different
mobilities in mung bean cell walls. Xyloglucans with a rigid
backbone comprised 27% of the xyloglucan; the rigid xyloglucans were assumed to be bound to cellulose microfibrils,
whereas the majority of the wall xyloglucan was less rigid and
surrounded by mobile polysaccharides. The authors calculated
that <8% of the cellulose surface was coated with xyloglucan.
Dick-Perez et al. (2011) reached a similar conclusion by use of
two- and three-dimensional ssNMR to detect cross-peaks (spin
transfer) between cellulose and matrix polysaccharides of
Arabidopsis cell walls. Extensive interactions (spin transfer) of
the cellulose surface chains were observed for pectins (rhamnose and galacturonic acid residues), but not for xyloglucan.
Cross-peaks between interior cellulose chains and xyloglucan
were seen, suggesting that some xyloglucan may be entrapped
within cellulose microfibrils. Because xyloglucan is the dominant hemicellulose in dicot primary cell walls, these studies concluded that cellulose surfaces in these growing walls are largely
free of tightly bound hemicellulose.
At first glance, this conclusion is perplexing. Where is all the
xyloglucan in the wall, if it is not coating cellulose? If there is
plenty of free cellulose surface in the wall, why doesn’t xyloglucan bind it? There is no dispute about the strong binding interaction between xyloglucan and cellulose. Studies in which
fluorescently labeled xyloglucan was used to stain growing
pea epicotyl cells observed extensive binding when endogenous
xyloglucan was previously removed from cell wall ‘ghosts’, but
found negligible staining of walls not pre-extracted to remove
188
xyloglucan (Hayashi et al. 1987). This result was interpreted to
mean that most of the cellulose was natively coated with xyloglucan. Does this contradict the NMR results?
The 2D and 3D ssNMR studies identified spin transfer between pectin and cellulose surfaces as well as between pectin
and xyloglucan (Dick-Perez et al. 2011, Dick-Perez et al. 2012,
Wang et al. 2012). Such transfer requires close proximity and
relatively slow molecular motions, which is indicative of
stable interactions but not necessarily tight binding. Why
were xyloglucan–cellulose interactions in low abundance,
given the well-documented affinity of xyloglucan for cellulose
surfaces? Two factors may limit xyloglucan–cellulose
interactions within the cell wall: xyloglucan selectivity for binding to the hydrophobic surface of cellulose and steric hindrance
by pectins. Regarding steric hindrance, we note that the cell
wall matrix is a crowded space. Pectins may physically block
access to cellulose surfaces and may also interact directly
with xyloglucan, limiting the ability of entangled xyloglucan
chains to align to the hydrophobic surface of cellulose. Pectin
content is approximately 3-fold that of xyloglucan in
Arabidopsis cell walls (White et al. 2014), which is typical of
many species. Hence crowding, limited diffusion and entanglement by the pectin network may kinetically limit xyloglucan
access to cellulose.
Additionally, as discussed above, the high-affinity binding
sites for xyloglucan binding may be restricted to the hydrophobic surface, which may be a small percentage of the total cellulose surface. In contrast, cellulose surface chains identified by
NMR include both the hydrophobic and hydrophilic surfaces.
Xyloglucan could occupy most of the hydrophobic surfaces
while still leaving the majority of cellulose surface chains
without rigidly bound xyloglucan. Xyloglucans bound to the
hydrophilic surfaces of cellulose are relatively mobile (Zhao
et al. 2014) and may be gradually removed by the trimming
action of XTH enzymes (Thompson and Fry 1997). Pectin binding, on the other hand, may be biased towards the hydrophilic
surfaces (but this point has evidently not been investigated).
Although pectin binds more weakly to cellulose than does
xyloglucan (Zykwinska et al. 2008, Gu and Catchmark 2013),
its greater abundance may result in preferential occupancy of
the hydrophilic surfaces with pectin chains while at the same
time sterically interfering with access by xyloglucan. An
additional possibility is that interactions with pectins draw
xyloglucans away from weak binding to the hydrophilic surface
of cellulose. This possibility could be tested by computational
approaches. The net result of all these effects would be stronger
NMR cross-peaks between cellulose and pectin (Wang et al.
2012).
The idea that there is sufficient xyloglucan to coat all cellulose surfaces assumes that xyloglucan has an extended shape,
not the coiled shape that it has in solution (Hayashi 1989, Levy
et al. 1997). What is the evidence that xyloglucan assumes a
fully extended form in the wall? This concept is partly based on
a questionable extrapolation of xyloglucan stiffness at the scale
of its persistence length to longer scales. In addition, EM images
of putative xyloglucan chains dried onto a mica surface show a
highly extended shape (Mccann et al. 1992), similar to that
Plant Cell Physiol. 56(2): 180–194 (2015) doi:10.1093/pcp/pcu204
observed with AFM (Morris et al. 2004). However, this extended
conformation is an effect of interaction with the mica surface;
conformation within the complex, hydrated cell wall may be
quite different. At this point there seems to be no compelling
evidence concerning xyloglucan conformation in the native
wall. The fact that 38% of the xyloglucan in cell walls of pea
stems could be solubilized by xyloglucan-specific endoglucanase (Pauly et al. 1999) suggests that much of the xyloglucan
has an enzyme-accessible, well-solvated coil conformation. This
percentage is probably an underestimate because the presence
of other wall polymers reduces enzyme access to xyloglucan.
Additional support for a coiled configuration comes from
ssNMR studies of spin diffusion from water to cell wall polysaccharides in which xyloglucans (and pectins) were seen to be
well hydrated (White et al. 2014). Xyloglucan tightly bound to
cellulose would be at least partially dehydrated. When pectins
were partially extracted by treatment with calcium chelator,
water mobility increased and spin transfer from water to xyloglucan and to cellulose was greatly slowed. The conclusion was
that the organization of water in the cell wall was controlled by
pectins, which were in intimate contact with both cellulose and
xyloglucan.
Microfibril entrapment
Several observations suggest that xyloglucans become
entrapped by microfibrils during cell wall assembly: (i) extraction of xyloglucan from cell walls requires high alkaline conditions that swell cellulose, converting it to cellulose II, whereas
xyloglucan that is bound to cellulose in vitro is solubilized with
much lower alkaline solutions (Hayashi 1989); (ii) some xyloglucan in pea epicotyl walls was not released until the walls
were completely digested by cellulase (Pauly et al. 1999); (iii)
cellulose crystallinity is higher in a xyloglucan-deficient line of
Arabidopsis mutant compared with the wild type (Dick-Perez
et al. 2011); and (iv) NMR cross-peaks were detected between
xyloglucan and internal cellulose chains (Dick-Perez et al. 2011).
Xyloglucan entrapment during microfibril formation (Atalla
et al. 1993) is a potential explanation of these observations.
Consistent with this idea, cellulose crystallinity was reduced
when cellulose-synthesizing cells of Gluconacetobacter xylinus
were cultured in the presence of xyloglucan (Atalla et al. 1993,
Whitney et al. 1995, Park et al. 2014).
Tethering microfibrils
Pauly et al. (1999) operationally defined three xyloglucan
domains in cell walls from etiolated pea stems: (1) a domain
that was solubilized by xyloglucan-specific endoglucanase
(XEG), releasing 38% of the xyloglucan; (2) a domain solubilized
by additional extraction with 4 M KOH (48% of the xyloglucan);
and (3) a residual domain (14%) that was solubilized only
upon digestion of the residue with cellulase. Domain (1) was
assumed to comprise loops and tethers between microfibrils,
forming the hypothetical load-bearing links between cellulose
microfibrils. Domain (2) was considered to be xyloglucan
strands tightly bound to cellulose surfaces, and domain (3)
was proposed to be entrapped within or between cellulose
microfibrils.
The inference that cell wall strength depends on xyloglucan
tethers between microfibrils was tested directly by treatment of
cell walls with substrate-specific endoglucanases in combination with biomechanical assessments of the cell wall (Park
and Cosgrove 2012b). Three endoglucanases from glycosyl
hydrolase family 12 (www.cazy.org) were used: XEG, which digests xyloglucan but not cellulose; CEG, which cuts unbranched
glucans (cellulose) but not xyloglucan, and CXEG (Cel12A),
which cuts both xyloglucan and cellulose. Of the three enzymes,
only CXEG loosened the cell wall, assessed by cell wall creep and
wall compliances. These results showed that XEG-accessible
xyloglucan did not strengthen the wall. A combined treatment
with CEG and XEG might be expected to mimic the effect of
CXEG, but paradoxically such treatment did not loosen the
wall. Resolution of this enigma led to the ‘biomechanical hotspot’ hypothesis: a small fraction of xyloglucan is interlaced
with cellulose chains, forming relatively inaccessible adhesion
zones linking two or more microfibrils (Fig. 3). Efficient digestion of such a structure would require an enzyme with both
xyloglucanase and cellulase activity. This requirement explains
why enzymes from the XTH family do not cause cell wall creep
(McQueen-Mason et al. 1993, Saladie et al. 2006) and why XTH
mutants exhibit negligible growth phenotype (Kaewthai et al.
2013, Hara et al. 2014). Genetic redundancy has sometimes
been invoked to account for the lack of phenotype in XTH
mutants, but Kaewthai et al. (2013) showed that XTH hydrolytic activity was completely eliminated in AtXTH31/AtXTH32
double knockout lines, yet they displayed negligible growth
phenotype.
Substantial wall loosening by CXEG was traced to the
digestion of a specific component comprising <1% of the xyloglucan in the wall, indicating that very limited sites may control
wall extensibility (Park and Cosgrove 2012b). The xxt1/xxt2
mutant lacked a creep response to CXEG, indicating that
xyloglucan was necessary for the formation of these biomechanical junctions (Park and Cosgrove 2012a). Moreover, CXEG
treatment solubilized some pectins as well as xyloglucan from
Arabidopsis walls and increased water–pectin interactions
(White et al. 2014), suggesting that pectins may be
connected to the biomechanical hotspots in some way, as yet
undefined.
Recent cell wall cartoons have depicted both xyloglucan and
pectins acting as parallel tethers between microfibrils
(Zykwinska et al. 2007, Dick-Perez et al. 2012). To evaluate
the relative significance of pectin and xyloglucan for wall extensibility, we used enzymatic and other biochemical treatments to loosen pectins or xyloglucans selectively in
Arabidopsis cell walls and measured the biomechanical consequences (Park and Cosgrove 2012a). The results indicated that
treatments targeting pectin had negligible effects compared
with those targeting the xyloglucan–cellulose junctions. The
effects of pectin loosening, however, were notably larger in
cell walls of the Arabidopsis xxt1/xxt2 line, indicating that pectins have a greater mechanical role when xyloglucan is absent.
One might expect a similar situation in celery parenchyma,
which contains only 2% xyloglucan (Thimm et al. 2002), but
this has not been tested. A recent mathematical model of
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Y. B. Park and D. J. Cosgrove | Xyloglucan interactions in growing cell walls
Fig. 3 Biomechanical hotspot model in which cellulose microfibril contacts are mediated by a subfraction of inaccessible xyloglucans. (A)
Cellulose microfibrils are depicted as rods with hydrophilic (yellow) and hydrophobic (blue) surfaces; extended xyloglucan chains (intense green
beads) are found in the cellulose–cellulose junctions while coiled regions act to disperse microfibrils. Pectins are not shown. (B) Computational
model of a cross-section of a junction between the hydrophobic surfaces of two cellulose microfibrils (blue) glued together through a monolayer
of xyloglucan. (C) Depiction of two lamellae in a primary cell wall. The cellulose is shown as an 18 chain rectangular model with yellow
hydrophilic surfaces and blue hydrophobic surfaces. Pectin chains (orange/red) fill much of the space between cellulose layers and make
extensive contracts with the hydrophilic surfaces of cellulose, whereas xyloglucans (green/blue) preferentially bind to the hydrophobic surfaces
of cellulose in limited regions and form coils between microfibrils. Image B adapted from Zhao et al. (2014), copyright Springer Verlag, and used
with permission.
growing cell walls proposed pectins to be a key determinant of
extensibility in yielding cell walls (Dyson et al. 2012), but experimental assessment of this idea is needed.
Control of Wall Extensibility at the Meso-Scale
The biomechanical hotspot hypothesis requires that cellulose
microfibrils come into close contact with each other at limited
mechanical junctions. This might be via a monolayer of xyloglucan binding the hydrophobic surfaces of two microfibrils
together (Fig. 3B, C). Such junctions are missing in common
depictions of cell walls where microfibrils are drawn as well
separated from each other and only linked via tethers. In contrast, recent AFM images of never-dried cell walls (Fig. 4) show
that cellulose microfibrils do come into close lateral contact
over short regions (Zhang et al. 2014). Because the walls in this
AFM study were imaged under water and were never dried,
artifactual aggregation that occurs during extraction and dehydration could be excluded. The idea of limited biomechanical
junctions is also consistent with a low density of a-expansin
190
binding sites in the cell wall (McQueen-Mason and Cosgrove
1995). In Fig. 4 we have pictorially represented the density of
expansin binding sites in the surface lamella of a primary cell
wall. They are limited to a small fraction of the cell wall, although the exact spatial locations are unknown. A recent study
used ssNMR to identify chemically expansin’s target in a complex cell wall (Wang et al. 2013). This is technically challenging
because the low density of binding sites means the concentration of expansin in a wall sample, even at expansin saturation, is
too low to detect by typical ssNMR methods. By combining 15N
and 13C labeling with a sensitization method called dynamic
nuclear polarization, it was possible to obtain an NMR spectrum of the polysaccharides within 13C spin-diffusion distance
(1 nm) of recombinant expansin proteins bound to the cell
wall. The NMR spectrum showed that the target was cellulose,
but with a structure slightly different from bulk cellulose and
with xyloglucan in close proximity. This characterization is remarkably similar to the hypothesized biomechanical hotspots,
based on a completely different approach.
These results lead to a more nuanced concept of the physical basis of cell wall extensibility. Conventional models have
Plant Cell Physiol. 56(2): 180–194 (2015) doi:10.1093/pcp/pcu204
require a suite of new approaches, combining the power of
genetic mutants with advances in imaging and labeling methods as well as physical characterization of the cell wall and
additional approaches yet to be devised.
Supplementary data
Supplementary data are available at PCP online.
Funding
This work was supported by the US Department of Energy,
Office of Science, Basic Energy Sciences as part of The Center
for LignoCellulose Structure and Formation, an Energy Frontier
Research Center under Award # DE-SC0001090.
Fig. 4 AFM image of the recently deposited surface of an onion epidermal outer cell wall. The red ellipses illustrate the average density of
a-expansin binding sites in the surface lamella, based on the binding
studies of McQueen-Mason and Cosgrove (1995). The arrows indicate
a few of the many regions where two microfibrils separate and come
into close contact. Image credit: the AFM image is reprinted from
Zhang et al. (2014), copyright Springer Verlag, and used with
permission.
envisaged the growing cell wall as a scaffold of cellulose microfibrils embedded in a polysaccharide matrix which acts to keep
microfibrils apart and to link them via load-bearing polysaccharide tethers. For >40 years the focus has been at the molecular scale of microfibril–matrix–microfibril interactions, i.e.
the nanometer scale, with the premise that the viscoelasticity of
the matrix determines wall extensibility. The new model shifts
focus to a larger scale and proposes that wall extensibility is
controlled at limited sites where microfibrils make close contact. When these sites are loosened by the action of expansins
or by CXEG-type enzymes, the microfibrils slide or separate,
perhaps at a rate that is influenced by the bulk viscoelasticity
of the microfibril–matrix network between biomechanical hotspots. Pectins are likely to play a major role in wall viscoelasticity. Figs. 3 and 4 graphically illustrate some of the key concepts
of primary cell wall structure that probably impact cell wall
extensibility.
Perspective and Future Directions
The ideas summarized above leave many questions for future
inquiry. How are the biomechanical junctions formed? What is
their structure and lifetime? Do non-crystalline or crystalline
regions of the cellulose microfibril take part in junction formation? Is there more than one kind of such site in the wall? What
is the relative significance of yielding by bulk wall viscoelasticity
vs. yielding at the biomechanical junctions for wall extensibility
and control of cell growth? The answers to these questions will
Disclosures
The authors have no conflicts of interest to declare.
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