Update on Mechanisms of Plant Cell Wall Biosynthesis
Update on Mechanisms of Plant Cell Wall
Biosynthesis: How Plants Make Cellulose and
Other (1/4)-b-D-Glycans1
Nicholas C. Carpita*
Department of Botany and Plant Pathology, and Bindley Bioscience Center, Purdue University, West
Lafayette, Indiana 47907–2054
The discovery of a gene that encodes a cotton
(Gossypium hirsutum) cellulose synthase (Pear et al.,
1996) revolutionized and invigorated the plant cell
wall community to find the genes that encode the
machinery of cell wall polysaccharide synthesis. The
landscape was framed by the completion of the genome sequence of Arabidopsis (Arabidopsis thaliana;
Arabidopsis Genome Initiative, 2000), which gave a
complete gene inventory for a model plant species, but
one with many genes yet to be annotated for function.
An estimated 10% of the plant genome, about 2,500
genes, is devoted to construction, dynamic architecture, sensing functions, and metabolism of the plant
cell wall. Based largely on prior discoveries of function
in prokaryotic organisms, most of the tentatively annotated genes are organized into gene families for
substrate generation, glycosyl transfer, targeting and
trafficking, cell wall rearrangement, and modification
by hydrolases, esterases, and lyases (Yong et al., 2005;
Penning et al., 2009). However, the biochemical activities of most enzymes involved in glycosyl transfer
within these families remain to be verified, and an
additional 40% of the genome encodes genes whose
functions are not known. As many of these proteins
contain secretory signal peptides (Arabidopsis Genome Initiative, 2000), it is reasonable to infer that
some have roles in cell wall construction.
The Arabidopsis cellulose synthase/cellulose synthaselike (CesA/Csl) gene superfamily, which includes 10 CesA
genes and 29 Csl genes in six distinct groups, was one
of the first large families to be described (Richmond
and Somerville, 2000), and comparative analyses of a
reference dicot, Arabidopsis, with a reference grass,
rice (Oryza sativa), revealed substantive differences in
family structures, adding two groups not seen in the
dicot genome (Hazen et al., 2002). Extension of these
annotations to compare all cell wall-related gene families of the grasses with those of the dicots reveals
some correlation of family structure with the differ1
This work supported by the Center for Direct Catalytic Conversion of Biomass to Biofuels, an Energy Frontier Research Center
funded by the U.S. Department of Energy, Office of Science, Office of
Basic Energy Sciences (award no. DE–SC0000997).
* E-mail [email protected].
www.plantphysiol.org/cgi/doi/10.1104/pp.110.163360
ences between plants with type I walls and those of the
grasses with type II walls (Fig. 1A; Penning et al.,
2009). For CesA genes and certain Csl genes, establishment of specific function for the synthases they encode comes from the analysis of mutants lacking a
particular function and, in some specific examples,
by heterologous expression. However, genetic approaches alone do not inform us about the biological
mechanism of synthesis. The knowledge gained from
molecular genetic approaches now needs to be augmented by biochemical and cell biological approaches
to achieve a greater understanding of proteins and
their interactions within a synthase complex, their
organization at membranes, and their dynamics. This
Update focuses on the biochemical mechanisms of the
synthesis of a single type of linkage, the (1/4)-b-D
linkage, in which one sugar is inverted nearly 180°
with respect to each neighboring sugar in the chain.
This linkage presents a unique steric problem for
processive catalysis that all living organisms have
solved but we are still struggling to understand.
This Update reviews our present state of knowledge
of the biochemical mechanisms of polysaccharide
synthesis, including some classic discoveries, and
presents an alternative hypothesis on the biochemical
mechanisms and organization of complexes involved
in synthase reactions that yield (1/4)-b-D linkages.
CELLULOSE SYNTHESIS
In flowering plants, cellulose is a para-crystalline
array of about two to three dozen (1/4)-b-D-glucan
chains. Microfibrils of 36 glucan chains have a theoretical diameter of 3.8 nm, but x-ray scattering and
NMR spectroscopy indicate that some microfibril diameters could be as small as 2.4 nm, or about two
dozen chains (Kennedy et al., 2007). The microfibrils
are synthesized at the plasma membrane by terminal
complexes of six-membered “particle rosettes” that
produce a single microfibril (Giddings et al., 1980;
Mueller and Brown, 1980). Thus, each of the six
components of the particle rosette is expected to
synthesize four to six of the glucan chains, and 24 to
36 chains are then assembled into a functional microfibril (Doblin et al., 2002). In freeze fracture, the par-
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Carpita
Figure 1. The CesA/Csl gene superfamily. A, Of the 10 Arabidopsis
CesA genes, at least three are coexpressed during primary wall formation, and mutations in each of them, AtCesA1 (RSW1; At4g32410),
AtCesA6 (PRC1; At5g64740), and AtCesA3 (CEV1 and ELI1;
At5g05170), result in cellulose deficiencies, indicating that each is
essential for cellulose synthesis. The irx mutants AtCesA8 (IRX1;
At4g18780), AtCesA7 (IRX3; At5g17420), and AtCesA4 (IRX5;
At5g44030) are deficient in cellulose synthesis specifically in secondary
walls. Seven additional subgroups were identified that are the likely
synthases for noncellulosic polysaccharides with backbones of (1/4)b-D-glycans. Whereas the CesA genes of Arabidopsis, rice, and maize
appear to be orthologous, the Csl genes are divergent between dicots
and grasses, species that make two distinct kinds of walls. From mutants
and heterologous expression studies, members of the CslA group encode
the synthases of (gluco)mannans, members of the CslC group are likely
to encode the glucan backbone of xyloglucans, and the rice- and maizeonly members of CslH and CslF encode the synthases of the mixedlinkage (1/3),(1/4)-b-D-glucans found only in grasses (after Penning
et al., 2009). B, Domain model and class-specific regions (CSRs) for
three CesAs known to function in primary cell wall cellulose synthesis.
Two ZnF domains (in yellow) are found in the N terminus before the first
membrane-spanning domain (in blue). Eight transmembrane helices,
two upstream and six downstream of the catalytic domain, are predicted
to interact to form a channel through which a single b-glucan chain is
secreted to the cell wall. The large central catalytic domain contains four
highly conserved “U motifs” of D, DxD, D, and QxxRW, important for
substrate binding and catalysis. Once thought to be a hypervariable
region (Pear et al., 1996), the class-specific regions are conserved among
orthologs of the same subclade and vary in the number of upstream
conserved Cys residues, the number of consecutive basic amino acids,
Lys and Arg, and the number of consecutive acidic amino acids, Asp and
Glu, downstream from the basic residues (after Carpita and Vergara,
1998; Vergara and Carpita, 2001).
ticle rosettes, found only on the P-face of the membrane, are about 25 nm in diameter, but this size
represents only the membrane-spanning and short
exterior domains (Fig. 2A). Hidden in surface views of
rosette structures in the plasma membrane, the much
larger catalytic domains of the cellulose synthases are
estimated to be 50 nm wide and extend 35 nm into the
cytoplasm (Bowling and Brown, 2008), a feature that
has escaped consideration in many published models
of the rosette structure (Fig. 2, B and C).
Cellulose synthase is an ancient enzyme (Nobles
et al., 2001), and cellulose synthase genes in green algae
are homologous to those of flowering plants (Roberts
et al., 2002). The deduced amino acid sequences of
CesAs share regions of similarity with the bacterial
CesA proteins, namely the four catalytic motifs containing the D, DxD, D, Q/RxxRW that are highly
conserved among those that synthesize several kinds
of (1/4)-b-D-glycans (Saxena et al., 1995). The higher
plant CesA genes are predicted to encode polypeptides
of about 110 kD, each with a large, cytoplasmic
N-terminal region containing zinc-finger (ZnF) domains,
and eight membrane spans sandwiching the four U
motifs of the catalytic domain (Fig. 1B; Delmer, 1999).
Further evidence for the functions of plant CesA
genes in cellulose synthesis came from Arabidopsis
mutants of three of the CesA genes involved in primary wall synthesis: the temperature-sensitive radial
swelling mutant rsw1 (AtCesA1; Arioli et al., 1998),
the dwarf-hypocotyl procuste mutant prc1 (AtCesA6;
Fagard et al., 2000), and a stunted root phenotype
with altered jasmonate and ethylene signaling (cev1)
and ectopic lignification (eli1) mutant alleles (AtCesA3;
Ellis and Turner, 2001; Caño-Delgado et al., 2003).
Despite coexpression in the same cells and an expectation of redundancy, cellulose synthesis is impaired in
each mutant. The same was observed with the irregular
xylem mutants irx1, irx3, and irx5, which display a
phenotype of collapsed mature xylem cells as a result
of lowered cellulose content during secondary cell
wall deposition (Taylor et al., 2000, 2003). A widely
accepted hypothesis is that the AtCesA1, AtCesA3,
and AtCesA6 proteins assemble to function in primary
wall cellulose synthesis, while the AtCesA4, AtCesA7,
and AtCesA8 proteins assemble to make secondary
wall cellulose (Fig. 1A), with each member of the trio
performing a nonredundant function in the complex
(Taylor, 2008). Lack of one CesA prevents incorporation of the other two into the plasma membrane
(Gardiner et al., 2003). However, at least some of the
subunits are potentially interchangeable, as inferred
by the dominant-negative inhibition of growth and
primary wall thickness caused by constitutive expression of a mutated fra5 (irx3 allele) transgene (Zhong
et al., 2003) and by the semi-dominant-negative phenotype observed in the heterozygous AtCesA3 mutant
(Daras et al., 2009). AtCesA1 is essential for cellulose
synthesis (Beeckman et al., 2002), whereas knockouts
of AtCesA3 (Ellis and Turner, 2001; Caño-Delgado
et al., 2003) and AtCes6 (Fagard et al., 2000) result in
partially impaired synthesis but not in total inhibition.
Desprez et al. (2007) indicated that the AtCesA2 and
AtCesA5 proteins have partially redundant functions
with AtCesA6.
A direct association of three distinct CesA polypeptides was demonstrated in vitro and by colocalization
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Synthesis of (1/4)-b-D-Glycans
rations subjected to native PAGE gave an 840-kD
complex and that null mutants, but not missense mutations, gave smaller 420-kD complexes (Wang et al.,
2008). Atanassov et al. (2009) affinity trapped a ladder
of complexes of CesA oligomers to about 700 to 730
kD. Consistent with the observations of Wang et al.
(2008), only smaller oligomeric complexes of two of
the CesAs are detected when the third is missing
(Atanassov et al., 2009). Such an association of CesAs
was indicated independently in yeast two-hybrid
studies (Timmers et al., 2009).
DOES SYNTHESIS OF EACH
(1/4)-b-D-GLUCAN CHAIN REQUIRE ONE OR
TWO CATALYTIC POLYPEPTIDES?
Figure 2. Particle rosette structures associated with cellulose synthesis
in angiosperms. A, Freeze-etch images of the P-face of the plasma
membrane showing clusters of rosettes associated with the developing
of secondary wall spiral thickenings of a Lepidium tracheary element
(from Herth, 1985). The inset shows the 6-fold symmetry of a single
particle rosette from a Zinnia tracheary element developing in vitro
(from C. Haigler, unpublished data, as seen in Delmer, 1999). A
substructure can be observed in each of the particles. In these freezeetch images, only the membrane-spanning domains and extracellular
loops of the CesA proteins can be observed. B, Cytoplasmic structure
(circled) underlying the rosettes in plasma membrane footprints (from
Bowling and Brown, 2008). These structures always are at the terminus
of a microfibril (arrow). Bar = 200 nm. C, A Markham rotational
analysis of one of these shows the reinforcement of hexagonal shape
with 60° rotational steps. All other angles of rotation cancel to circular
(Bowling and Brown, 2008).
in vivo by Taylor et al. (2003). Domain-swap experiments with wild-type and mutant AtCesA1 and
AtCesA3 proteins in their respective mutants resulted
in dominant-positive and dominant-negative effects,
consistent with both catalytic and C-terminal domains
being important for function (Wang et al., 2006). Direct
interactions of three distinct CesA polypeptides in
vivo were shown by bimolecular fluorescence complementation (Desprez et al., 2007). Although some
complementary pairs gave stronger fluorescence
than others, both homodimers and heterodimers of
AtCesA1, AtCesA3, and AtCesA6 are inferred. Wang
et al. (2008) used pull-down experiments similar to
those of Taylor et al. (2003) to show that these three
primary wall CesA proteins interact. Furthermore,
they showed that Triton-soluble microsomal prepa-
After over four decades of study, the biochemical
mechanism by which cellulose is made remains a
mystery (Delmer, 1999; Saxena and Brown, 2005;
Somerville, 2006; Guerriero et al., 2010), with only a
few reports of cellulose synthesis in vitro with isolated
membranes (Kudlicka and Brown, 1997; Lai-Kee-Him
et al., 2002). For both cellulose and the related (1/4)b-D-glycan, chitin, synthesis proceeds by the attachment of glucosyl residues to the nonreducing terminus
of the acceptor glucan chain (Koyama et al., 1997; Imai
et al., 2003). The simplest hypothesis is that each CesA
polypeptide synthesizes a single glucan chain. In the
Delmer (1999) model, the eight membrane spans form
a channel through which a single glucan chain is
extruded (Fig. 3A). This mode of synthesis comes with
a very big steric problem for synthesis. To make a (1/
4)-b-D linkage means that each glucosyl residue is
turned 180° with respect to each neighbor. Thus, the
O-4 position of nonreducing terminal sugar of the
acceptor chain is displaced several angstroms upon
addition of each successive unit (Fig. 3B). For the next
glycosyl transfer to occur, the site of catalysis must
move several angstroms within the protein, the acceptor chain must swivel 180°, or the catalytic or acid-base
amino acids must toggle between two forms to account for the displacement. To overcome this steric
problem, several models have proposed that two sites
or modes of glycosyl transfer reside within the catalytic complex, so that disaccharide units are added
iteratively (Carpita et al., 1996; Koyama et al., 1997;
Carpita and Vergara, 1998; Buckeridge et al., 1999, 2001;
Saxena et al., 2001) or that two polypeptides associate
to form two opposing catalytic sites (Buckeridge et al.,
2001; Vergara and Carpita, 2001). In either model,
glycobiosyl units, or any even-numbered oligomeric
units, are added to the nonreducing end to ensure that
the (1/4)-b-D linkages are strictly preserved without
inversion of substrate, active site, or terminus of the
growing chain (Fig. 3C).
Despite the rationale for a two-site model of catalysis, biochemical evidence from other types of
polysaccharide synthases indicate that a single
polypeptide is sufficient. Hyaluronan (HA) is an un-
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Carpita
Figure 3. Models for cellulose synthase and the steric problem of making a (1/4)-b-D-glycosyl linkage. A, The first model of
conformation for a single CesA protein subunit was proposed by Delmer (1999). Each CesA subunit must interact with other such
subunits to form the synthase complex. The ZnFs, plant-specific conserved region (P-CR), and class-specific region (CSR) are
potential interaction sites (figure modified from Delmer, 1999). B, The steric problem of synthesis is illustrated in top view and
end view. Addition of a single glycosyl residue in a (1/4)-b-D linkage without rotation of the end of the chain or the active site of
the synthase would result in movement of the O-4 several angstroms. C, The conceptual solution to the steric problem is a
catalytic dimer of simultaneous glycosyl transfer to form a cellobiosyl residue to the O-4 position of the terminal glucosyl residue
of the chain. Synthesis of an even number of units always maintains the acceptor position as the O-4 position as the chain is
extruded. In the catalytic dimer model, if one of the sites is damaged, then the point of attachment becomes the O-3 position,
which would maintain the point of attachment as the O-3, hence producing callose thereafter (after Buckeridge et al., 1999,
2001).
branched polysaccharide composed of repeating
units of (1/3)-b-D-GlcNAc and (1/4)-b-D-GlcUA
(DeAngelis and Weigel, 1994). HA synthases exist in
three distinct classes, with class I containing integral
membrane proteins to transport the HA across the
membrane (Fig. 4). Bacterial and mammalian HA
synthases have been shown to contain both transferase
activities in a single polypeptide (DeAngelis and
Weigel, 1994; Yoshida et al., 2000; Williams et al.,
2006). Such a finding argues that the synthesis of a
single HA polymer requires only a single polypeptide.
The crystal structure of a nonprocessive family 2
glycosyltransferase (GTs) sharing sequence similarity with a portion of the catalytic domain of a CesA,
the Bacillus subtilis SpsA synthase, provided the first
conformation of the active site and the role of the
aspartyl residues in the positioning of the uridinyl
group of a UDP-sugar (Fig. 5A; Charnock and Davies,
1999; http://www.pdb.org/pdb/explore/explore.do?
structureId=1QGS). Charnock and colleagues (2001)
argued that only a single site for a nucleotide-sugar
substrate is accommodated within a single polypeptide
of SpsA.
synthesis of a repeating (1/4)-b-D-glucosyl linkage
of the cellulose glucan chains. In fact, two recent
studies demonstrate unequivocally that at least some
HA synthases and homologs of SpsA synthase form
dimers. An identical structure to the SpsA synthase is
the 3BCV polypeptide from Bacteroides fragilis, which
is also predicted to contain a single substrate-binding
site. In contrast to SpsA synthase, the 3BCV protein
crystallizes as a dimer, and each monomer possesses
a bound UDP (Fig. 5B; http://www.pdb.org/pdb/
explore/explore.do?structureId=3BCV). The dimerization occurs through the C-terminal regions, which
appear to be flexible and, for this reason, were deleted
A CATALYTIC DIMER HYPOTHESIS
From the studies of the class I HA synthases and the
SpsA crystal structure, it is inferred that a single
polypeptide alone has all the features needed for
synthesis. However, these features still do not address
mechanistically the fundamental steric problem of
Figure 4. Models of HA synthases. A, Class I synthases. B, Class II
synthases (after Weigel and DeAngelis, 2007).
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Synthesis of (1/4)-b-D-Glycans
Figure 5. Crystal structures of type 2 glycosyl transferases. A, The SpsA synthase crystallizes as a monomer with a single binding site for UDP (Charnock
and Davies, 1999; http://www.pdb.org/pdb/explore/
explore.do?structureId=1QGS). B, The B. fragilis
SpsA homologous polypeptide crystallizes as a
dimer (http://www.pdb.org/pdb/explore/explore.do?
structureId=3BCV), each with a single binding domain for UDP. C, Each member of the crystal dimer
E. coli chondroitin polymerase has two UDP-GlcAor UDP-binding domains (http://www.pdb.org/pdb/
explore/explore.do?structureId=2Z86).
from the crystal structure of the SpsA synthase
(Charnock and Davies, 1999). This, to our knowledge,
is the first direct evidence by crystal structure of a
homodimer formed by GT2 proteins, but even more
complicated structures are also found. For example,
an Escherichia coli strain K4 chondroitin polymerase
contains two “Rossmann fold-like” domains within
a single polypeptide, each binding a UDP-GlcA or
UDP, and it also crystallizes as a dimer, giving a total of
four nucleotide-binding domains (Fig. 5C; http://
www.pdb.org/pdb/explore/explore.do?structureId=
2Z86). Although the fit is not exceptionally good, the
CesA catalytic domain threads through the E. coli
chondroitin polymerase best of all for known crystal
structures of glycosyl transferases involving nucleotide sugars (D. Kihara, personal communication), and
it is consistent with the suggestion by Brown and
Saxena (2000) and Saxena et al. (2001) of a conformation within which the catalytic domain of a single
CesA would allow the synthesis of cellobiose units of
the chain within a single polypeptide.
The class II synthases contain two different types of
GT2 modules but not the membrane-spanning domains (Fig. 4). One type of HA synthase possesses two
repeats of the UDP-Glc and acceptor-binding domains, so the synthesis of the characteristic disaccharide of HA by a single synthase is rationalized (Jing
and DeAngelis, 2000). However, a direct interaction of
two synthases is inferred for HA synthesis to explain
the finding that host cells harboring constructs in
which each site is independently disrupted are still
able to make HA (Jing and DeAngelis, 2000; Weigel
and DeAngelis, 2007). Because the class I HA synthases both have activities on a single peptide does not
preclude the possibility that the formation of homodimers of single isoforms of HA synthase is necessary
for function, which, like the chrondroitin polymerase,
would give four nucleotide sugar-binding sites per
dimer.
Solving the steric problem aside, one must also ask if
a channel of eight membrane spans proposed for the
cellulose synthase is of sufficient size to extrude a (1/
4)-b-D-glucan chain. The question about sufficient
channel size was raised also with respect to the HA
synthases by Weigel and DeAngelis (2007), who suggested that certain phospholipids required for activity
possibly integrate with the membrane spans to widen
the channel for extrusion. However, whether lipid
interactions with a small number of domains would be
significant is still in question. Callose synthases are
about twice the size of CesAs and contain 16 membrane spans (i.e. double those of a CesA; Hong et al.,
2001). Plasma membrane hexose and maltose transporters of prokaryotes and eukaryotes are homologous (Maiden et al., 1987), and virtually all of them
contain a minimum of 11 to as many as 18 transmembrane spans per functional unit (Reifenberger et al.,
1995; Pao et al., 1998; Sherson et al., 2000; Klepek et al.,
2010).
The sensitivity of detergent-solubilized CesA complexes to dithiothreitol suggested to Atanassov et al.
(2009) that disulfide bonds are involved in the coupling into larger complexes. Other features of the
protein outside the region of catalysis, such as the
ZnFs, which show high similarity to RING-finger
domains that bind zinc in a “cross-brace” manner
(Freemont, 2000), might function in the organization of
these into the larger rosette structure. Kurek et al.
(2002) proposed that CesAs are coupled through the
ZnF domain in a redox-dependent manner, constituting the first step in the clustering of CesAs into
rosettes. Moreover, the discovery that a thioredoxinlike protein associates with the CesA ZnF domain in
a yeast two-hybrid screen led to the suggestion
that reduction of the domains by oxidoreductases
returns the CesAs to monomeric forms, which are directed to the ubiquitin-dependent turnover pathway
(Kurek et al., 2002). The experimental herbicide CGA
325#615 blocks crystallization of the b-D-glucan chains
into cellulose microfibrils, phenocopying the rsw1
swollen root tip (Peng et al., 2001). This phenotype
can be abrogated completely in the presence of hydro-
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Carpita
gen peroxide, suggesting that the inhibitor blocks
rosette assembly by enzymatic oxidation (Kurek
et al., 2002).
If ZnF domains of two CesAs couple as part of
the recruitment into rosette particles, the question
remaining is how all the others interact to form a
complete complex. Although not discussed specifically, the study by Kurek et al. (2002) presented data
that full-length CesA proteins formed tetramers and
even higher ordered pairings, while the ZnF domains
were limited to coupling of a single pair. Timmers
et al. (2009) showed that heterodimer interactions
indicated by yeast two-hybrid analysis do not require
the ZnF domains. Taken together, these data provide
evidence that domains other than the ZnFs of the
CesA participate in coupling reactions if two to three
dozen CesAs or more are aggregated to form a rosette
complex.
Comparison of CesA sequences suggests potential
heterodimeric interaction domains within the catalytic
domain. The initial scarcity of CesA protein sequences
and the apparent variability within the so-called
“hypervariable region” led to the assumption that
this region was probably not essential in catalysis
(Pear et al., 1996). However, it is now understood that
these regions are well conserved across grass and dicot
species with a distinct subclade structure. Potential
protein-protein interactions through subdomains of
this region containing conserved Cys residues, clusters
of consecutive basic Lys and Arg residues, and clusters
of acidic Asp and Glu residues form the basis of a
class-specific region (Fig. 1B; Vergara and Carpita,
2001).
To test the catalytic dimer hypothesis, we expressed
fusion proteins containing only the catalytic domain of
Arabidopsis and maize (Zea mays) CesAs with affinity
tags and observed that dimers and higher order aggregates collapse reversibly to monomeric forms by
thiol-reducing agents (C. Rayon, A. Olek, L. Paul, and
S. Ghosh, unpublished data). Because the ZnF was
absent in these constructs, dimerization must occur
through thiol-sensitive sequences in the catalytic
domain. Such an interaction of CesAs to form
homodimers or heterodimers solves the three basic
problems of the single polypeptide-single polymer
conundrum: (1) the steric problem is solved by coordinate synthesis and attachment of cellobiose units
instead of monomers, preserving the integrity of the
O-4 site of attachment at the nonreducing terminus of
the chain; (2) a channel of 16 membrane-spanning
domains is consistent with sugar transport and callose
extrusion; and (3) the interaction produces two ZnF
domains for recruitment of the catalytic dimer into
rosette particles (Fig. 6). An exciting prospect is that
conservation of space would be maintained if CesAs
turn out to have a structure like the chondroitin
polymerase dimers and function like class II HA
synthases, because a CesA catalytic dimer with four
nucleotide-binding domains would be capable of
generating two (1/4)-b-D-glucan chains instead of
Figure 6. A catalytic dimer hypothesis for cellulose synthase. A, A
catalytic dimer model of two CesAs to form a complex that synthesizes
a single (1/4)-b-D-glucan chain. Homodimerization or heterodimerization of CesAs gives mirrored active sites that generate cellobiosyl
units, which are then attached to the nonreducing end of the extruded
glucan chain. Dimerization also results in a channel composed of 16
membrane-spanning domains, equivalent to that of callose synthase
and consistent with eukaryotic monosaccharide transporters. B, Dimerization results in two ZnF domains that are now able to couple two
neighbors instead of just one. C, Six such complexes interact to
constitute one particle of the six-particle rosette.
just one. Further experiments are needed to establish
preferred heterodimer interactions, the stoichiometry of UDP-Glc binding, and the role of the ZnF in
recruitment of the catalytic dimers into larger complexes.
THE BIOLOGICAL SYNTHESIS OF CELLULOSE
Cellulose synthase has a half-life of less than 30 min,
remarkably short for a membrane protein (Jacob-Wilk
et al., 2006). Assembly of rosettes occurs in the Golgi
stacks, and they must be continually secreted to the
plasma membrane to maintain cellulose synthesis
(Haigler and Brown, 1986). Additional proteins are
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Synthesis of (1/4)-b-D-Glycans
suspected to be necessary for the formation of primers
of polymer synthesis, metabolic channeling of substrates, crystallization of the chains, and termination of
chains (Doblin et al., 2002; Somerville, 2006; Guerriero
et al., 2010). Furthermore, a proteomics survey of
plasma membrane proteins shows that certain CesAs
are phosphorylated at several locations within the
catalytic and N-terminal domains (Nühse et al., 2004).
Modification of some potential phosphorylation sites
with amino acids that either prevent (Ala) or mimic
(Glu) phosphorylation has multiple effects that reduce
either synthesis rates or interactions with the microtubule cytoskeleton independently (Chen et al., 2010).
In affinity-labeling experiments with [32P]UDP-Glc,
an 84-kD polypeptide was found to be associated with
a plasma membrane fraction containing the highest
activity of callose synthase (Delmer et al., 1991) and
subsequently was identified as Suc synthase (SuSy).
Confirmation of plasma membrane association was
made immunocytochemically (Amor et al., 1995), and
Delmer and Amor (1995) proposed that the association
of SuSy represented a UDP-Glc delivery mechanism to
cellulose synthase. b-Glucan microfibrils are synthesized from Suc and UDP on immobilized tobacco
(Nicotiana tabacum) plasma membrane sheets (Hirai
et al., 1998). More recently, SuSy was immunologically
associated with CesA proteins in the rosette structures
(Fujii et al., 2010), strengthening the idea of an association of SuSy directly with cellulose synthases for
metabolic channeling of Glc through a localized pool
of UDP-Glc. However, a quadruple mutant that eliminates all detectable SuSy in vegetative tissue does not
impair cellulose synthesis (Barratt et al., 2009). Overexpression of SuSy in developing vascular tissue of
transgenic poplar yields small but significant increases
in cellulose content (Coleman et al., 2009). Taken
together, SuSy does not appear to be required for
cellulose synthesis but may enhance rates by concentrating substrate at the site of synthesis.
Peng et al. (2002) provided evidence that sitosterolcellodextrins synthesized from sitosterol-b-glycoside
serve as primers of glucan chain initiation, with the
KORRIGAN glucanohydrolase trimming the sitosterol
from the growing chain. DeBolt and colleagues (2009)
question this role, as they found that double mutants
of two major sterol-b-glucoside synthases result in
severe defects in cuticle formation but not in cellulose
synthesis. However, the sterol-glucosides are substantially reduced in the double mutant but not entirely
eliminated, leaving open the question.
Outside the Golgi stacks, a membrane compartment
also containing KORRIGAN (Robert et al., 2005) might
represent a dynamic factory associated with both the
microtubule network and the plasma membrane that
aligns and directs the cellulose synthase complex and
coordinate that function with the deposition of the many
polysaccharides directed to it from packaged Golgi
vesicles. Bimolecular fluorescence complementation
techniques, as shown for CesA interactions (Desprez
et al., 2007), can give important clues to selected
participants in protein-protein interactions within a
complex in vivo. Fluorescence tagging has also allowed visualization of the movement of cellulose
synthase complexes at the plasma membrane (Paredez
et al., 2006; Wightman and Turner, 2008; Gutierrez
et al., 2009). These studies also established the dynamics of the relationships with the cortical microtubule
network in real time. As reviewed by Baskin (2001)
and Szymanski and Cosgrove (2009), these studies
bring resolution to ideas on the alignment of cortical
microtubules and cellulose microfibrils generated long
ago through observations with inhibitors (Green, 1962)
and in the electron microscope (Ledbetter and Porter,
1963). Improvements in imaging tools are still needed
that permit visualization of the delivery of Golgiderived vesicles to the sites of cellulose synthesis, and
much progress has already been made along these
lines (Held et al., 2008; Konopka and Bednarek, 2008;
Crowell et al., 2009).
THE SYNTHESIS OF NONCELLULOSIC
POLYSACCHARIDES WITH
(1/4)-b-D-GLYCAN BACKBONES
Because of the same conserved domains of nucleotide-sugar binding and catalysis as those encoding
CesAs, subfamilies of Csl genes were predicted to
encode the synthases of noncellulosic polymers with
(1/4)-b-D-glycan backbones, primarily (gluco)mannans and galacto(gluco)mannans, xyloglucans, glucuronoarabinoxylans (GAX), and the grass-specific (1/
3),(1/4)-b-D-glucans (Delmer, 1999; Richmond and
Somerville, 2000; Hazen et al., 2002). For the most part,
this has turned out to be true, but GAXs are a clear
exception. There is still an incomplete knowledge of
most of the gene products and interactions among
them to make specific b-D-glycans, even among families where at least one member has a confirmed
glycosyl transferase activity. There is also a growing
disconnection between the classic studies on the synthesis of these polysaccharides in vitro and the discovery of genes encoding the machinery that warrants
a revisit.
CslF AND CslH: MIXED-LINKAGE
(1/3),(1/4)-b-D-GLUCAN SYNTHASE IS THE
TOPOLOGICAL EQUIVALENT OF
CELLULOSE SYNTHASE
The mixed-linkage (1/3),(1/4)-b-D-glucan is
made in the grasses (Poales; Carpita, 1996; Buckeridge
et al., 2004), certain lichens (Wood et al., 1994), and
Equisetum (Fry et al., 2008; Sørensen et al., 2008), but
differences in the distribution of their cellodextrin
oligomers indicate that they probably arose by convergent evolution of synthases. For the grasses, this
glucan is not a random mixture of (1/3)-b-D- and
(1/4)-b-D-glucosyl linkages but is composed primarily of cellotriosyl and cellotetraosyl units in a ratio of
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Carpita
about 2.5:1 connected by single (1/3)-b-D linkages
(Wood et al., 1994). Upon cleavage with a Trichoderma
cellulase, smaller amounts of higher cellodextrin series
are observed, with the odd-numbered cellodextrin
2-fold higher in abundance than the next even-numbered unit in the series. The synthesis of the (1/3),
(1/4)-b-D-glucan has been demonstrated in vitro
with isolated, intact Golgi membranes and UDP-Glc
(Gibeaut and Carpita, 1993; Buckeridge et al., 1999,
2001; Urbanowicz et al., 2004). Whereas micromolar
concentrations of UDP-[14C]Glc result in much shorter
oligomers and polymers enriched in cellotetraosyl
units rather than cellotriosyl units (Buckeridge et al.,
1999), much larger polysaccharides enriched in cellotriosyl units are observed with higher concentrations
of substrate (Buckeridge et al., 1999, 2001). The ratios
of cellotriosyl-cellotetraosyl and cellopentosyl-cellohexaosyl are increased proportionally with substrate
concentrations higher than 250 mM (Buckeridge et al.,
1999), indicating that the mechanism of synthesis of
the odd-numbered cellodextrin unit is fundamentally
different from synthesis of the even-numbered units.
Proteolysis protection assays show further that the
active site of catalysis is on the outward-facing Golgi
membrane (Urbanowicz et al., 2004). Golgi membranes treated with proteinase K specifically lost their
ability to make the odd-numbered cellodextrin units,
whereas the synthesis of the cellotetraosyl and higher
order even-numbered units was unaffected. Again,
loss of the ability to make cellotriosyl units is correlated with significant loss in size of the (1/3),(1/4)b-D-glucan product (Urbanowicz et al., 2004). We
proposed a similar catalytic dimer model wherein
even-numbered units are synthesized by core cellulose
synthase-like proteins and the odd-numbered units
arise by an additional GT that has yet to be identified
(Buckeridge et al., 2001, 2004).
Limited proteolysis and detergent reconstitution
experiments with the mixed-linkage (1/3),(1/4)-bD-glucan of grass species provides kinetic evidence for
three sites of glycosyl transfer within the catalytic
domain: two from the cellulose synthase-like core
domain and a third, separable activity (Urbanowicz
et al., 2004). (1/3),(1/4)-b-D-Glucan is the topological equivalent of cellulose synthase at the Golgi
membrane. Limited proteolysis or detergent treatment
causes loss of the ability to make the diagnostic oddnumbered cellotriose units for synthesis without affecting the ability to generate the even-numbered
cellotetraosyl unit (Urbanowicz et al., 2004).
As the topologic equivalent of cellulose synthase at
the Golgi membrane, the (1/3),(1/4)-b-D-glucan
synthase shares another feature with cellulose synthase. When its resident membranes are damaged,
cellulose synthase (Delmer, 1977) and the (1/3),(1/
4)-b-D-glucan synthase (Buckeridge et al., 2001) “default” to synthesis of the (1/3)-b-D-glucan, callose,
possibly by disruption of the complete active site to a
single glycosyl transferase activity (Buckeridge et al.,
2001; Urbanowicz et al., 2004). Loss of the cellobiosyl-
generating system to a single site of glycosyl transfer
would thereafter make only callose (Buckeridge et al.,
2001), whose synthesis does not require turning the
catalytic site or acceptor 180° (Fig. 3B). While it can be
argued that membrane disruption activates callose
synthase in vitro in plasma membrane preparations
of all angiosperms (Nishimura et al., 2003), only Golgi
membranes from grasses, the only angiosperms that
make the (1/3),(1/4)-b-D-glucan, make callose when
damaged (Gibeaut and Carpita, 1993; Buckeridge et al.,
1999). The most direct evidence for the default synthesis of callose from a damaged cellulose synthase comes
from the experiments of Blanton et al. (2000), who
showed that isolated membranes of a cellulose synthase mutant of the cellular slime mold Dictyostelium
discoideum also lost the ability to make callose in vitro.
Whereas some cellulose is made in vitro with wild-type
membrane preparations, callose linkages predominate.
Because cellulose synthase is a single gene in Dictyostelium, loss of the ability to make (1/3)-b-D-glucan in
vitro as well as (1/4)-b-D-glucan in membranes from
the mutant strongly suggests that the single polypeptide is responsible for both activities.
Two groups of Csl genes, CslF and CslH, which are
found only in grasses (Hazen et al., 2002), have been
shown to catalyze (1/3),(1/4)-b-D-glucan biosynthesis. Heterologous expression of a rice CslF in
Arabidopsis, a species that does not make (1/3),
(1/4)-b-D-glucan, results in small amounts of the b-Dglucan in the cell walls (Burton et al., 2006). However,
considerably greater amounts of the (1/3),(1/4)-bD-glucan result when a CslH is coexpressed with CslF,
suggesting a synergistic role for both CslH and CslF in
the synthesis of the polysaccharide and that a catalytic
heterodimer enhances the activity (Doblin et al., 2009).
If an accessory glycosyl transferase is necessary to
make the odd-numbered cellodextrin unit, then Arabidopsis must produce a related isoform. This finding
of concerted action by two distinct group members
highlights the possibility that synthases of other crosslinking glycans might be encoded by Csl genes of
different groups.
CslA: MANNAN AND
GLUCOMANNAN BIOSYNTHESIS
One of the first cell wall polysaccharides to be
synthesized in vitro was glucomannan. An early conclusion that GDP-Glc is the substrate for (1/4)-b-D
linkages of cellulose and that UDP-Glc is the substrate
for (1/3)-b-D-glucans (Chambers and Elbein, 1970)
had already been disproven, yet GDP-Glc is still listed
erroneously as the substrate for cellulose synthesis on
most wall charts of biochemical pathways. Kinetic
evidence obtained with cotton fiber cells cultured in
vitro showed unequivocally that UDP-Glc is the substrate for cellulose synthesis (Carpita and Delmer,
1981). Addition of both GDP-Glc and GDP-Man to
membrane preparations resulted in marked stimula-
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Synthesis of (1/4)-b-D-Glycans
tion of incorporation into a glucomannan product
(Elbein and Hassid, 1966; Piro et al., 1993).
The knowledge of GDP-nucleotide sugars as substrates was instrumental in the discovery that a CslA
gene encodes a mannan synthase by expression profiling of guar (Cyamopsis tetragonolobus) seed development, a species that accumulates large amounts of
galactomannan as a cell wall storage carbohydrate
(Dhugga et al., 2004). Liepman et al. (2005, 2007)
confirmed that at least four members of the CslA
group function in mannan and/or glucomannan synthesis. However, mixed substrates of GDP-Glc and
GDP-Man in the heterologous expression system
(Liepman et al., 2007) do not give the marked enhancement of glucomannan synthesis long ago observed in vitro (Elbein and Hassid, 1966). While the
CesA genes appear orthologous across several species,
the Csl genes are not (Penning et al., 2009). In fact, the
CslA group is resolved into three subgroups that either
are Arabidopsis dominated, grass dominated, or mixed.
The CslA members defined as mannan synthase genes
(Liepman et al., 2007) fall into both the Arabidopsisdominated and the mixed subgroups (Penning et al.,
2009). The functions of these other subgroup members
of the grasses need to be defined.
CslC: XYLOGLUCAN BIOSYNTHESIS
Xyloglucans were among the first complex cell wall
polysaccharides whose synthesis was demonstrated in
vitro. Early studies showed that labeled sugars from
UDP-Glc and UDP-Xyl are incorporated into several
polysaccharides using microsomal membranes and
were later refined by isolation of Golgi membranes
(Ray et al., 1969; Ray, 1980). Small amounts of xyloglucan-like oligomers with the characteristic a- D -Xyl(1/6)- D -glucosyl unit, isoprimeverose, are made
with small amounts of UDP-Glc and UDP-Xyl, but
Gordon and Maclachlan (1989) found that when concentrations of each nucleotide-sugar are increased to
millimolar levels, large polymers containing the characteristic heptasaccharide XXXG (for nomenclature,
see Table I) unit structure are synthesized. The tetraglucosyl unit of the xyloglucan backbone and the
precisely repeated three xylosyl units added to make
the XXXG structure are consistent with an even-numbered cellobiose unit synthase reaction for the glucan
backbone. Even in the structural variant of solanaceous xyloglucan, where just two xylosyl units are
added, a tetraglucosyl unit backbone is preserved by
the replacement of the third xylosyl group with an
acetate (Sims et al., 1996). An apparent exception is the
ability of certain tree legumes, such as jatobá (Hymenaea courbaril), to make XXXXG units in addition to
XXXG (Buckeridge et al., 2000). Curiously, partial
digestion of this polymer with a Trichoderma cellulase,
which cleaves only at unbranched positions, yields
octomer, nonamer, and decamer backbone oligomers
whose ratios predict that the polymer consists of 4-55-4 frameworks separated by variable amounts of
XXXG, rather than a random distribution of XXXXG
and XXXG units (Tiné et al., 2006). This is an intriguing
result for two reasons: (1) the 4-5-5-4 framework of
these types of xyloglucans preserves the even-numbered unit symmetry of the backbone; and (2) to make
such a framework, as many as 18 glucosyl residues
might be contained within the complex in order to be
“read” properly to preserve the unit structure, making
the complex much larger than expected.
The topology of xyloglucan synthesis at the Golgi
membrane is still uncertain. Several lines of evidence
suggest that synthesis relies on transporters for UDPGlc for backbone synthesis of (1/4)-b-D-glucans in
vitro (Orellana, 2005), and UDP-GlcA is transported
to provide UDP-Xyl for transfer within the lumen
(Hayashi et al., 1988). Lerouxel et al. (2006) proposed
that synthesis of the backbone is the topological
equivalent of cellulose synthase, but the addition of
all subtending sugars of Xyl, Gal, and Fuc are within
the lumen of the Golgi.
Based on heterologous expression in Pichia, Cocuron
et al. (2007) provide evidence that CslC genes encode
the synthases of the xyloglucan backbone. Although
Pichia is unable to make UDP-Xyl, coexpression of the
xylosyl transferase with CslC is sufficient to induce the
extension of (1/4)-b-D-glucan chains, indicating that
a close interaction of these proteins might stabilize the
synthase to allow extension of the backbone. A complication to unequivocal annotation of function of this
subfamily is the finding of a CslC at the plasma
membrane instead of the Golgi membrane (Dwivany
et al., 2009). Three members of the GT34 family are
established as the xylosyl transferases involved in
xyloglucan synthesis, and these Golgi-resident proteins are predicted to face the lumen (Cavalier et al.,
2008; Zabotina et al., 2008).
Xyloglucan is decorated in various ways in a species-specific manner (Hoffman et al., 2005; Peña et al.,
2008). Most are primarily galactosylated, with a characteristic a-L-Fuc-(1/2)-b-D-Gal-(1/2)-a-D-Xyl trisaccharide extension (Bauer et al., 1973), but others,
such as those of solanaceous species, have a truncated
unit structure substituted with a-L-Ara-(1/2)-a-DXyl extensions instead of Gal (Sims et al., 1996), and
the Asteridae and Oleales species have mixtures of
these two forms of xyloglucan substitution (Hoffman
et al., 2005). Furthermore, the xyloglucans synthesized in the Golgi are modified in ways that make
them structurally different from those that are assembled onto the cellulose microfibrils in the wall
(Obel et al., 2009).
The purification of a xyloglucan-specific fucosyl
transferase led to the discovery of a GT37 gene encoding it (Perrin et al., 1999). While the synthase complex
must involve a close interaction between the glucan
synthases and the xylosyl transferases that decorate it
(Cavalier et al., 2008), the association of the galactosyl
and fucosyl transferases might be more transient.
Transferases extracted from the membrane are able
to add Gal from UDP-Gal (Madson et al., 2003) and
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Carpita
Table I. Six possible xyloglucan oligomers are produced by digestion
with a Trichoderma endoglucanase
Because of characteristic side group construction of these complex
glucans, a single-letter code was devised to denote the nonreducing
terminal sugar of the side chain, with other constituents understood
(Fry et al., 1993). Thus, XXXG signifies a tetraglucosyl backbone with
three residues bearing a xylosyl side group. For many angiosperms, this
heptasaccharide is decorated further with variable amounts of Gal at
the two Xyl residues closer to the reducing end, and if Gal is added to
the first Xyl residue, a Fuc residue is usually added. In addition to the
glucan backbone synthase, at least three types of glycosyl transferases,
and as many as six, are needed to construct all the side groups.
Oligomer
Notation
Xyl
Xyl
Glc 2 Glc 2 Glc 2 Glc
Xyl
XXXG
Xyl
Xyl
Glc 2 Glc 2 Glc 2 Glc
Xyl
Gal
Gal
Xyl
Xyl
Glc 2 Glc 2 Glc 2 Glc
Xyl
Fuc
Gal
Xyl
Xyl
Glc 2 Glc 2 Glc 2 Glc
Xyl
Gal
Xyl
Xyl
Glc 2 Glc 2 Glc 2 Glc
Xyl
Gal
Fuc
Gal
Xyl
Xyl
Glc 2 Glc 2 Glc 2 Glc
Xyl
Gal
XLXG
XXLG
XXFG
XLLG
XLFG
Fuc from GDP-Fuc (Perrin et al., 1999; Vanzin et al.,
2002) to exogenous xyloglucan in vitro.
XYLAN BIOSYNTHESIS
The synthesis of grass (1/4)-b-D-xylans from UDPXyl with microsomal membranes was first demonstrated by Bailey and Hassid (1966). Cooperative
action of two nucleotide-sugar substrates, in this instance UDP-Xyl and UDP-GlcA, resulted in the synthesis of (1/4)-b-D-xylans with subtending GlcA
units (Waldron and Brett, 1983; Baydoun et al., 1989).
Similar studies have shown that membrane preparations from grasses and mixtures of nucleotide sugars
made GAXs, in part employing a nascent C-4 epimerase that interconverts UDP-Xyl and UDP-Ara (Porchia
and Scheller, 2000; Kuroyama and Tsumuraya, 2001;
Porchia et al., 2002; Zeng et al., 2008).
Apart from a hint that the AtCslD5 may be involved
(Bernal et al., 2007) in the synthesis of (1/4)-b-Dxylans, it has yet to be directly demonstrated that any
Csl gene plays a role. In fact, informatics approaches
yield non-Csl genes as more likely candidates for encoding the machinery for xylan synthesis (Mitchell
et al., 2007), and several mutants with deficiencies in
normal xylan synthesis, such as parvus (Lao et al., 2003;
Lee et al., 2007), irx8 (Brown et al., 2005), irx7/fra8
(Zhong et al., 2005; Brown et al., 2007), irx9 and irx14
(Brown et al., 2007), and irx10 and irx10-L (Brown et al.,
2009), do not include a member of the Csl gene family.
(1/4)-b-D-Glucuronoxylan (GX) synthesis appears
to involve a complex initiation sequence. Peña et al.
(2007) discovered that collapsed xylem mutants deficient in xylan, irx8 and fra8, were essentially devoid of
a complex tetrasaccharide, b-D-Xyl-(1/3)-a-L-Rha(1/2)-a-D-GalA-(1/4)-D-Xyl, located at the reducing
end of the xylan polymer, whereas an irx9 mutant also
severely deficient in xylan contained an overabundance of the tetrasaccharide. The presence or absence
of this tetrasaccharide greatly affected the size distribution of the xylans. In irx8 and fra8, a broader
distribution is observed, with some polymers longer
than observed in the wild type. Xylans of irx9 have
short chains, with nearly all of them containing the
tetrasaccharide (Peña et al., 2007). These results suggested a model whereby short chains of (1/4)-b-Dxylan are primed by the tetrasaccharide, and these are
spliced, cleaving the primer, to make the long polysaccharides (York and O’Neill, 2008).
The IRX8 (GAUT12) and PARVUS (GATL1) genes
encode members of the GT8 group C, and IRX7/FRA8
genes encode members of the GT47 group E, that
synthesize the primer tetrasaccharide, whereas IRX9
and IRX14 encode members of GT43 that are likely to
encode the synthases of the (1/4)-b-D-xylan oligomeric backbones that are stitched together by a yet
unidentified glycosyl transferase (Brown et al., 2007;
Peña et al., 2007). GT8 family members encode retaining-type transferases, and the GT47 members encode
inverting-type transferases with respect to the anomeric linkages formed compared with the anomeric linkage in the nucleotide sugar, so GT8 members make a-D
linkages, whereas the GT47 members make b-D or a-L
linkages. While an a-D-GalA is found in the tetrasaccharide, (1/2)-a-D-GlcA (4-O-Me-GlcA) side groups
are also attached at precise intervals along the xylan
chain (Nishitani and Nevins, 1991) and a general
blockwise synthesis of six consecutive branched xylosyl residues in grass xylans is observed (Carpita and
Whittern, 1986). IRX10 and IRX10-L appear to encode
xylosyl transferases also from GT47, but double mutants of these genes have greatly reduced GlcA substitutions along the shorter chains (Brown et al., 2009).
It will be interesting to determine if the addition of
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Synthesis of (1/4)-b-D-Glycans
these GlcA side groups plays a role as an attachment
or recognition point where short xylan chains are
grafted together to make a long chain, a suggestion
foretold by the original work on the cooperative action
of UDP-GlcA and UDP-Xyl in the in vitro synthesis of
glucuronoxylan (Waldron and Brett, 1983; Baydoun
et al., 1989).
York and O’Neill (2008) take a broader perspective
of xylan synthesis and have suggested that reducing
end addition should not be ruled out. In their model, a
type of alternating “pendulum” mechanism is proposed for the introduction of the (1/4)-b-D-xylan
linkages, a mechanism analogous to the dimer synthesis described here for (1/4)-b-D linkages in general. We are just learning the identities of the GlcA and
Araf transferases of the GX and GAX polymers and the
distinctions between the GX devoid of Ara that is
abundant in the secondary xylem and the GAX with its
rich Ara substitution that is the major polymer of the
primary cell walls of grasses (Scheller and Ulvskov,
2010). Recently, proteomic approaches of isolated GAX
synthase complexes from wheat membranes demonstrated a close interaction of GT43 and GT47 family
members with a GT75 UDP-Ara mutase, an enzyme
that interconverts the UDP-arabinopyranose and
UDP-arabinofuranose conformations required for incorporation of the latter into the polysaccharide (Zeng
et al., 2010). These kinds of studies represent the path
forward to identify all the components of synthase
complexes whose activities are preserved in vitro.
CONCLUDING REMARKS
While the plant cell wall community is making
steady progress in defining biochemical functions of
backbone synthases and the glycosyl transferases that
decorate them, the biochemical details of these large,
coordinated complexes are still unknown. Fading
from memory are a great many foundation studies of
polysaccharide synthesis in vitro that give insights to
the differences between an isolated activity and the
function of a complex as a whole.
The major differences between the behaviors of protein complexes in vitro and in vivo are compounded
by the topology of the synthases and associated glycosyl transferases at the plasma membrane and Golgi
membranes, across which both pH gradients and
electrical potentials are generated. Early studies with
excised cotton fibers cultured in vitro showed that
resealing of the plasma membrane was essential to
reconstitute cellulose synthesis, and evidence provided that regeneration of the electrical potential
rather than the pH gradient led to that was critical to
the preservation of synthesis (Carpita and Delmer,
1980). This work was extended to show that artificial
potentials stimulate (1/3)-b-D-glucan synthesis in
vitro (Bacic and Delmer, 1981). In Gluconacetobacter,
rates of cellulose synthesis could be modulated directly through adjustment of the electrical potential in
living cells under conditions that did not impair
metabolism (Delmer et al., 1982). Such gradients exist
across other compartments, and, in contrast to cellulose synthesis at the plasma membrane, it is maintenance of a pH gradient that prolongs the synthesis of
the mixed-linkage (1/3),(1/4)-b-D-glucan in vitro in
isolated maize Golgi membranes (Gibeaut and Carpita,
1993). The physiological and biochemical bases for the
effect of these potentials and gradients on polysaccharide synthesis are still not understood. They do
serve to illustrate that, beyond the biochemical mechanism of synthesis, technologies that preserve not only
the protein complexes but also their cellular context
need to be developed to truly understand the synthesis
of macromolecules across membrane surfaces.
ACKNOWLEDGMENTS
I thank Maureen McCann (Purdue University) and Peter Ulvskov (University of Copenhagen) for their review of the manuscript and their many
helpful suggestions. I also thank Daisuke Kihara (Purdue University) for his
contributions to the discussion on protein structure and modeling. The
artwork in Figure 6 is by Pamela Burroff-Murr (Purdue University).
Received July 24, 2010; accepted November 2, 2010; published November 4,
2010.
LITERATURE CITED
Amor Y, Haigler CH, Johnson S, Wainscott M, Delmer DP (1995) A
membrane-associated form of sucrose synthase and its potential role in
synthesis of cellulose and callose in plants. Proc Natl Acad Sci USA 92:
9353–9357
Arabidopsis Genome Initiative (2000) Analysis of the genome sequence of
the flowering plant Arabidopsis thaliana. Nature 408: 796–815
Arioli T, Peng LC, Betzner AS, Burn J, Wittke W, Herth W, Camilleri C,
Höfte H, Plazinski J, Birch R, et al (1998) Molecular analysis of cellulose
biosynthesis in Arabidopsis. Science 279: 717–720
Atanassov II, Pittman JK, Turner SR (2009) Elucidating the mechanisms of
assembly and subunit interaction of the cellulose synthase complex of
Arabidopsis secondary cell walls. J Biol Chem 284: 3833–3841
Bacic A, Delmer DP (1981) Stimulation of membrane-associated polysaccharide synthetases by a membrane potential in developing cotton
fibers. Planta 152: 346–351
Bailey RW, Hassid WZ (1966) Xylan synthesis from uridine-diphosphateD -xylose by particulate preparations from immature corncobs. Proc Natl
Acad Sci USA 56: 1586–1593
Barratt DHP, Derbyshire P, Findlay K, Pike M, Wellner N, Lunn J, Feil R,
Simpson C, Maule AJ, Smith AM (2009) Normal growth of Arabidopsis
requires cytosolic invertase but not sucrose synthase. Proc Natl Acad Sci
USA 106: 13124–13129
Baskin TI (2001) On the alignment of cellulose microfibrils by cortical
microtubules: a review and a model. Protoplasma 215: 150–171
Bauer WD, Talmadge KW, Keegstra K, Albersheim P (1973) The structure
of plant cell walls. II. The hemicellulose of the walls of suspensioncultured sycamore cells. Plant Physiol 51: 174–187
Baydoun EAH, Waldron KW, Brett CT (1989) The interaction of xylosyltransferase and glucuronyltransferase involved in glucuronoxylan synthesis in pea (Pisum sativum) epicotyls. Biochem J 257: 853–858
Beeckman T, Przemeck GK, Stamatiou G, Lau R, Terryn N, De Rycke R,
Inzé D, Berleth T (2002) Genetic complexity of cellulose synthase A gene
function in Arabidopsis embryogenesis. Plant Physiol 130: 1883–1893
Bernal AJ, Jensen JK, Harholt J, Sørensen S, Moller I, Blaukopf C,
Johansen B, de Lotto R, Pauly M, Scheller HV, et al (2007) Disruption of
ATCSLD5 results in reduced growth, reduced xylan and homogalacturonan synthase activity and altered xylan occurrence in Arabidopsis.
Plant J 52: 791–802
Plant Physiol. Vol. 155, 2011
181
Downloaded from on July 31, 2017 - Published by www.plantphysiol.org
Copyright © 2011 American Society of Plant Biologists. All rights reserved.
Carpita
Blanton RL, Fuller D, Iranfar N, Grimson MJ, Loomis WF (2000) The
cellulose synthase gene of Dictyostelium. Proc Natl Acad Sci USA 97:
2391–2396
Bowling AJ, Brown RM Jr (2008) The cytoplasmic domain of the cellulosesynthesizing complex in vascular plants. Protoplasma 233: 115–127
Brown DM, Goubet F, Wong VW, Goodacre R, Stephens E, Dupree
P, Turner SR (2007) Comparison of five xylan synthesis mutants
reveals new insight into the mechanisms of xylan synthesis. Plant J 52:
1154–1168
Brown DM, Zeef LAH, Ellis J, Goodacre R, Turner SR (2005) Identification
of novel genes in Arabidopsis involved in secondary cell wall formation
using expression profiling and reverse genetics. Plant Cell 17: 2281–2295
Brown DM, Zhang Z, Stephens E, Dupree P, Turner SR (2009) Characterization of IRX10 and IRX10-like reveals an essential role in glucuronoxylan biosynthesis in Arabidopsis. Plant J 57: 732–746
Brown RM Jr, Saxena IM (2000) Cellulose biosynthesis: a model for
understanding the assembly of biopolymers. Plant Physiol Biochem
38: 57–67
Buckeridge MS, dos Santos HP, Tiné MAS (2000) Mobilisation of storage
cell wall polysaccharides in seeds. Plant Physiol Biochem 38: 141–152
Buckeridge MS, Rayon C, Vergara CE, Carpita NC (2004) Mixed linkage
(1/3)(1/4)-b-D-glucans of grasses. Cereal Chem 81: 115–127
Buckeridge MS, Vergara CE, Carpita NC (1999) The mechanism of synthesis of a mixed-linkage (1/3), (1/4)b-D-glucan in maize: evidence
for multiple sites of glucosyl transfer in the synthase complex. Plant
Physiol 120: 1105–1116
Buckeridge MS, Vergara CE, Carpita NC (2001) Insight into multi-site
mechanisms of glycosyl transfer in (1/4)b-D-glycans provided by the
cereal mixed-linkage (1/3),(1/4)b-D-glucan synthase. Phytochemistry 57: 1045–1053
Burton RA, Wilson SM, Hrmova M, Harvey AJ, Shirley NJ, Medhurst A,
Stone BA, Newbigin EJ, Bacic A, Fincher GB (2006) Cellulose synthaselike CslF genes mediate the synthesis of cell wall (1,3;1,4)-b-D-glucans.
Science 311: 1940–1942
Caño-Delgado A, Penfield S, Smith C, Catley M, Bevan M (2003) Reduced
cellulose synthesis invokes lignification and defense responses in
Arabidopsis thaliana. Plant J 34: 351–362
Carpita N, McCann M, Griffing LR (1996) The plant extracellular matrix:
news from the cell’s frontier. Plant Cell 8: 1451–1463
Carpita NC (1996) Structure and biogenesis of the cell walls of grasses.
Annu Rev Plant Physiol Plant Mol Biol 47: 445–476
Carpita NC, Delmer DP (1980) Protection of cellulose synthesis in detached cotton fibers by polyethylene glycol. Plant Physiol 66: 911–916
Carpita NC, Delmer DP (1981) Concentration and metabolic turnover of
UDP-glucose in developing cotton fibers. J Biol Chem 256: 308–315
Carpita NC, Vergara CE (1998) A recipe for cellulose. Science 279: 672–673
Carpita NC, Whittern D (1986) A highly substituted glucuronoarabinoxylan from developing maize coleoptiles. Carbohydr Res 146: 129–140
Cavalier DM, Lerouxel O, Neumetzler L, Yamauchi K, Reinecke A,
Freshour G, Zabotina OA, Hahn MG, Burgert I, Pauly M, et al (2008)
Disrupting two Arabidopsis thaliana xylosyltransferase genes results in
plants deficient in xyloglucan, a major primary cell wall component.
Plant Cell 20: 1519–1537
Chambers J, Elbein AD (1970) Biosynthesis of glucans in mung bean
seedlings: formation of b-(1,4)-glucans from GDP-glucose and b-(1,3)glucans from UDP-glucose. Arch Biochem Biophys 138: 620–631
Charnock SJ, Davies GJ (1999) Structure of the nucleotide-diphosphosugar transferase, SpsA from Bacillus subtilis, in native and nucleotidecomplexed forms. Biochemistry 38: 6380–6385
Charnock SJ, Henrissat B, Davies GJ (2001) Three-dimensional structures
of UDP-sugar glycosyltransferases illuminate the biosynthesis of plant
polysaccharides. Plant Physiol 125: 527–531
Chen S, Ehrhardt DW, Somerville CR (2010) Mutations of cellulose
synthase (CESA1) phosphorylation sites modulate anisotropic cell expansion and bidirectional mobility of cellulose synthase. Proc Natl Acad
Sci USA 107: 17188–17193
Cocuron JC, Lerouxel O, Drakakaki G, Alonso AP, Liepman AH, Keegstra
K, Raikhel N, Wilkerson CG (2007) A gene from the cellulose synthaselike C family encodes a b-1,4 glucan synthase. Proc Natl Acad Sci USA
104: 8550–8555
Coleman HD, Yan J, Mansfield SD (2009) Sucrose synthase affects carbon
partitioning to increase cellulose production and altered cell wall
ultrastructure. Proc Natl Acad Sci USA 106: 13118–13123
Crowell EF, Bischoff V, Desprez T, Rolland A, Stierhof Y-D, Schumacher
K, Gonneau M, Höfte H, Vernhettes S (2009) Pausing of Golgi bodies on
microtubules regulates secretion of cellulose synthase complexes in
Arabidopsis. Plant Cell 21: 1141–1154
Daras G, Rigas S, Penning B, Milioni D, McCann MC, Carpita NC,
Fasseas C, Hatzopoulos P (2009) The thanatos mutation in Arabidopsis
thaliana cellulose synthase 3 (AtCesA3) has a dominant-negative effect
on cellulose synthesis and plant growth. New Phytol 184: 114–126
DeAngelis PL, Weigel PH (1994) Immunochemical confirmation of the
primary structure of streptococcal hyaluronan synthase and synthesis of
high molecular weight product by the recombinant enzyme. Biochemistry 33: 9033–9039
DeBolt S, Scheible WR, Schrick K, Auer M, Beisson F, Bischoff V,
Bouvier-Navé P, Carroll A, Hematy K, Li Y, et al (2009) Mutations in
UDP-glucose:sterol glucosyltransferase in Arabidopsis cause transparent testa phenotype and suberization defect in seeds. Plant Physiol 151:
78–87
Delmer DP (1977) Biosynthesis of cellulose and other plant cell wall
polysaccharides. Rec Adv Phytochem 11: 105–153
Delmer DP (1999) Cellulose biosynthesis: exciting times for a difficult field
of study. Annu Rev Plant Physiol Plant Mol Biol 50: 245–276
Delmer DP, Amor Y (1995) Cellulose biosynthesis. Plant Cell 7: 987–1000
Delmer DP, Benziman M, Padan E (1982) Requirement for a membrane
potential for cellulose synthesis in intact cells of Acetobacter xylinum.
Proc Natl Acad Sci USA 79: 5282–5286
Delmer DP, Solomon M, Read SM (1991) Direct photolabeling with [32P]
UDP-glucose for identification of a subunit of cotton fiber callose
synthase. Plant Physiol 95: 556–563
Desprez T, Juraniec M, Crowell EF, Jouy H, Pochylova Z, Parcy F, Höfte H,
Gonneau M, Vernhettes S (2007) Organization of cellulose synthase
complexes involved in primary cell wall synthesis in Arabidopsis
thaliana. Proc Natl Acad Sci USA 104: 15572–15577
Dhugga KS, Barreiro R, Whitten B, Stecca K, Hazebroek J, Randhawa GS,
Dolan M, Kinney AJ, Tomes D, Nichols S, et al (2004) Guar seed
b-mannan synthase is a member of the cellulose synthase super gene
family. Science 303: 363–366
Doblin MS, Kurek I, Jacob-Wilk D, Delmer DP (2002) Cellulose biosynthesis in plants: from genes to rosettes. Plant Cell Physiol 43:
1407–1420
Doblin MS, Pettolino FA, Wilson SM, Campbell R, Burton RA, Fincher
GB, Newbigin E, Bacic A (2009) A barley cellulose synthase-like CSLH
gene mediates (1,3;1,4)-b-D-glucan synthesis in transgenic Arabidopsis.
Proc Natl Acad Sci USA 106: 5996–6001
Dwivany FM, Yulia D, Burton RA, Shirley NJ, Wilson SM, Fincher GB,
Bacic A, Newbigin E, Doblin MS (2009) The CELLULOSE-SYNTHASE
LIKE C (CSLC) family of barley includes members that are integral
membrane proteins targeted to the plasma membrane. Mol Plant 2:
1025–1039
Elbein AD, Hassid WZ (1966) The enzymatic synthesis of a glucomannan.
Biochem Biophys Res Commun 23: 311–318
Ellis C, Turner JG (2001) The Arabidopsis mutant cev1 has constitutively
active jasmonate and ethylene signal pathways and enhanced resistance
to pathogens. Plant Cell 13: 1025–1033
Fagard M, Desnos T, Desprez T, Goubet F, Refregier G, Mouille G,
McCann M, Rayon C, Vernhettes S, Höfte H (2000) PROCUSTE1
encodes a cellulose synthase required for normal cell elongation specifically in roots and dark-grown hypocotyls of Arabidopsis. Plant Cell
12: 2409–2424
Freemont PS (2000) Ubiquitination: RING for destruction? Curr Biol 10:
R84–R87
Fry SC, Nesselrode BHWA, Miller JG, Mewburn BR (2008) Mixed-linkage
(1/3,1/4)-b-D-glucan is a major hemicellulose of Equisetum (horsetail) cell walls. New Phytol 179: 104–115
Fry SC, York WS, Albersheim P, Darvill A, Hayashi T, Joseleau JP, Kato Y,
Lorences EP, Maclachlan GA, McNeil M, et al (1993) An unambiguous
nomenclature for xyloglucan-derived oligosaccharides. Physiol Plant
89: 1–3
Fujii S, Hayashi T, Mizuno K (2010) Sucrose synthase is an integral
component of the cellulose synthesis machinery. Plant Cell Physiol 51:
294–301
Gardiner JC, Taylor NG, Turner SR (2003) Control of cellulose synthase
complex localization in developing xylem. Plant Cell 15: 1740–1748
Gibeaut DM, Carpita NC (1993) Synthesis of (1/3), (1/4)-b-D-glucan in
182
Plant Physiol. Vol. 155, 2011
Downloaded from on July 31, 2017 - Published by www.plantphysiol.org
Copyright © 2011 American Society of Plant Biologists. All rights reserved.
Synthesis of (1/4)-b-D-Glycans
the Golgi apparatus of maize coleoptiles. Proc Natl Acad Sci USA 90:
3850–3854
Giddings TH Jr, Brower DL, Staehelin LA (1980) Visualization of particle
complexes in the plasma membrane of Micrasterias denticulata associated
with the formation of cellulose fibrils in primary and secondary cell
walls. J Cell Biol 84: 327–339
Gordon R, Maclachlan G (1989) Incorporation of UDP-[ 14C]glucose into
xyloglucan by pea membranes. Plant Physiol 91: 373–378
Green PB (1962) Mechanism for plant cellular morphogenesis. Science 138:
1404–1405
Guerriero G, Fugelstad J, Bulone V (2010) What do we really know about
cellulose biosynthesis in higher plants? J Integr Plant Biol 52: 161–175
Gutierrez R, Lindeboom JJ, Paredez AR, Emons AMC, Ehrhardt DW
(2009) Arabidopsis cortical microtubules position cellulose synthase
delivery to the plasma membrane and interact with cellulose synthase
trafficking compartments. Nat Cell Biol 11: 797–806
Haigler CH, Brown RM Jr (1986) Transport of rosettes from the Golgi
apparatus to the plasma membrane in isolated mesophyll cells of Zinnia
elegans during differentiation to tracheary elements in suspension culture. Protoplasma 134: 111–120
Hayashi T, Koyama T, Matsuda K (1988) Formation of UDP-xylose and
xyloglucan in soybean Golgi membranes. Plant Physiol 87: 341–345
Hazen SP, Scott-Craig JS, Walton JD (2002) Cellulose synthase-like genes
of rice. Plant Physiol 128: 336–340
Held MA, Boulaflous A, Brandizzi F (2008) Advances in fluorescent
protein-based imaging for the analysis of plant endomembranes. Plant
Physiol 147: 1469–1481
Herth W (1985) Plasma-membrane rosettes involved in localized wall
thickening during xylem vessel formation of Lepidium sativum L. Planta
164: 12–21
Hirai N, Sonobe S, Hayashi T (1998) In situ synthesis of b-glucan microfibrils on tobacco plasma membrane sheets. Proc Natl Acad Sci USA 95:
15102–15106
Hoffman M, Jia ZH, Peña MJ, Cash M, Harper A, Blackburn AR II,
Darvill A, York WS (2005) Structural analysis of xyloglucans in the
primary cell walls of plants in the subclass Asteridae. Carbohydr Res
340: 1826–1840
Hong Z, Delauney AJ, Verma DPS (2001) A cell plate-specific callose
synthase and its interaction with phragmoplastin. Plant Cell 13: 755–768
Imai T, Watanabe T, Yui T, Sugiyama J (2003) The directionality of chitin
biosynthesis: a revisit. Biochem J 374: 755–760
Jacob-Wilk D, Kurek I, Hogan P, Delmer DP (2006) The cotton fiber zincbinding domain of cellulose synthase A1 from Gossypium hirsutum
displays rapid turnover in vitro and in vivo. Proc Natl Acad Sci USA 103:
12191–12196
Jing W, DeAngelis PL (2000) Dissection of the two transferase activities of
the Pasteurella multocida hyaluronan synthase: two active sites exist in
one polypeptide. Glycobiology 10: 883–889
Kennedy CJ, Cameron GJ, Šturcová A, Apperley DC, Altaner C, Wess TJ,
Jarvis MC (2007) Microfibril diameter in celery collenchyma cellulose:
x-ray scattering and NMR evidence. Cellulose 14: 235–246
Klepek YS, Volke M, Konrad KR, Wippel K, Hoth S, Hedrich R, Sauer N
(2010) Arabidopsis thaliana POLYOL/MONOSACCHARIDE TRANSPORTERS 1 and 2: fructose and xylitol/H+ symporters in pollen and
young xylem cells. J Exp Bot 61: 537–550
Konopka CA, Bednarek SY (2008) Variable-angle epifluorescence microscopy: a new way to look at protein dynamics in the plant cell cortex.
Plant J 53: 186–196
Koyama M, Helbert W, Imai T, Sugiyama J, Henrissat B (1997) Parallel-up
structure evidences the molecular directionality during biosynthesis of
bacterial cellulose. Proc Natl Acad Sci USA 94: 9091–9095
Kudlicka K, Brown RM Jr (1997) Cellulose and callose biosynthesis in
higher plants. I. Solubilization and separation of (1/3)- and (1/4)b-glucan synthase activities from mung bean. Plant Physiol 115:
643–656
Kurek I, Kawagoe Y, Jacob-Wilk D, Doblin M, Delmer D (2002) Dimerization of cotton fiber cellulose synthase catalytic subunits occurs via
oxidation of the zinc-binding domains. Proc Natl Acad Sci USA 99:
11109–11114
Kuroyama H, Tsumuraya Y (2001) A xylosyltransferase that synthesizes
b-(1/4)-xylans in wheat (Triticum aestivum L.) seedlings. Planta 213:
231–240
Lai-Kee-Him J, Chanzy H, Müller M, Putaux JL, Imai T, Bulone V (2002)
In vitro versus in vivo cellulose microfibrils from plant primary wall
synthases: structural differences. J Biol Chem 277: 36931–36939
Lao NT, Long D, Kiang S, Coupland G, Shoue DA, Carpita NC, Kavanagh
TA (2003) Mutation of a family 8 glycosyltransferase gene alters cell wall
carbohydrate composition and causes a humidity-sensitive semi-sterile
dwarf phenotype in Arabidopsis. Plant Mol Biol 53: 647–661
Ledbetter MC, Porter KR (1963) A “microtubule” in plant cell fine structure. J Cell Biol 19: 239–250
Lee CH, Zhong RQ, Richardson EA, Himmelsbach DS, McPhail BT, Ye
ZH (2007) The PARVUS gene is expressed in cells undergoing secondary
wall thickening and is essential for glucuronoxylan biosynthesis. Plant
Cell Physiol 48: 1659–1672
Lerouxel O, Cavalier DM, Liepman AH, Keegstra K (2006) Biosynthesis of
plant cell wall polysaccharides: a complex process. Curr Opin Plant Biol
9: 621–630
Liepman AH, Nairn CJ, Willats WGT, Sørensen I, Roberts AW, Keegstra
K (2007) Functional genomic analysis supports conservation of
function among cellulose synthase-like A gene family members
and suggests diverse roles of mannans in plants. Plant Physiol 143:
1881–1893
Liepman AH, Wilkerson CG, Keegstra K (2005) Expression of cellulose
synthase-like (Csl) genes in insect cells reveals that CslA family members
encode mannan synthases. Proc Natl Acad Sci USA 102: 2221–2226
Madson M, Dunand C, Li X, Verma R, Vanzin GF, Caplan J, Shoue DA,
Carpita NC, Reiter WD (2003) The MUR3 gene of Arabidopsis encodes a
xyloglucan galactosyltransferase that is evolutionarily related to animal
exostosins. Plant Cell 15: 1662–1670
Maiden MCJ, Davis EO, Baldwin SA, Moore DCM, Henderson PJF (1987)
Mammalian and bacterial sugar transport proteins are homologous.
Nature 325: 641–643
Mitchell RAC, Dupree P, Shewry PR (2007) A novel bioinformatics
approach identifies candidate genes for the synthesis and feruloylation
of arabinoxylan. Plant Physiol 144: 43–53
Mueller SC, Brown RM Jr (1980) Evidence for an intramembrane component associated with a cellulose microfibril-synthesizing complex in
higher plants. J Cell Biol 84: 315–326
Nishimura MT, Stein M, Hou BH, Vogel JP, Edwards H, Somerville SC
(2003) Loss of a callose synthase results in salicylic acid-dependent
disease resistance. Science 301: 969–972
Nishitani K, Nevins DJ (1991) Glucuronoxylan xylanohydrolase: a unique
xylanase with the requirement for appendant glucuronosyl units. J Biol
Chem 266: 6539–6543
Nobles DR, Romanovicz DK, Brown RM Jr (2001) Cellulose in cyanobacteria: origin of vascular plant cellulose synthase? Plant Physiol 127:
529–542
Nühse TS, Stensballe A, Jensen ON, Peck SC (2004) Phosphoproteomics
of the Arabidopsis plasma membrane and a new phosphorylation site
database. Plant Cell 16: 2394–2405
Obel N, Erben V, Schwarz T, Kühnel S, Fodor A, Pauly M (2009)
Microanalysis of plant cell wall polysaccharides. Mol Plant 2: 922–932
Orellana A (2005) Biosynthesis of non-cellulosic polysaccharides in the
Golgi apparatus: topological considerations. Plant Biosyst 139: 42–45
Pao SS, Paulsen IT, Saier MH Jr (1998) Major facilitator superfamily.
Microbiol Mol Biol Rev 62: 1–34
Paredez AR, Somerville CR, Ehrhardt DW (2006) Visualization of cellulose
synthase demonstrates functional association with microtubules. Science 312: 1491–1495
Pear JR, Kawagoe Y, Schreckengost WE, Delmer DP, Stalker DM (1996)
Higher plants contain homologs of the bacterial celA genes encoding the
catalytic subunit of cellulose synthase. Proc Natl Acad Sci USA 93:
12637–12642
Peña MJ, Darvill AG, Eberhard S, York WS, O’Neill MA (2008) Moss and
liverwort xyloglucans contain galacturonic acid and are structurally
distinct from the xyloglucans synthesized by hornworts and vascular
plants. Glycobiology 18: 891–904
Peña MJ, Zhong R, Zhou GK, Richardson EA, O’Neill MA, Darvill AG,
York WS, Ye ZH (2007) Arabidopsis irregular xylem8 and irregular xylem9:
implications for the complexity of glucuronoxylan biosynthesis. Plant
Cell 19: 549–563
Peng L, Kawagoe Y, Hogan P, Delmer D (2002) Sitosterol-b-glucoside as
primer for cellulose synthesis in plants. Science 295: 147–150
Peng LC, Xiang F, Roberts E, Kawagoe Y, Greve LC, Kreuz K, Delmer DP
(2001) The experimental herbicide CGA 325#615 inhibits synthesis of
Plant Physiol. Vol. 155, 2011
183
Downloaded from on July 31, 2017 - Published by www.plantphysiol.org
Copyright © 2011 American Society of Plant Biologists. All rights reserved.
Carpita
crystalline cellulose and causes accumulation of non-crystalline b-1,4glucan associated with CesA protein. Plant Physiol 126: 981–992
Penning BW, Hunter CT III, Tayengwa R, Eveland AL, Dugard CK, Olek
AT, Vermerris W, Koch KE, McCarty DR, Davis MF, et al (2009) Genetic
resources for maize cell wall biology. Plant Physiol 151: 1703–1728
Perrin RM, DeRocher AE, Bar-Peled M, Zeng W, Norambuena L, Orellana
A, Raikhel NV, Keegstra K (1999) Xyloglucan fucosyltransferase, an
enzyme involved in plant cell wall biosynthesis. Science 284: 1976–1979
Piro G, Zuppa A, Dalessandro G, Northcote DH (1993) Glucomannan
synthesis in pea epicotyls: the mannose and glucose transferases. Planta
190: 206–220
Porchia AC, Scheller HV (2000) Arabinoxylan biosynthesis: identification
and partial characterization of b-1,4-xylosyltransferase from wheat.
Physiol Plant 110: 350–356
Porchia AC, Sørensen SO, Scheller HV (2002) Arabinoxylan biosynthesis
in wheat: characterization of arabinosyltransferase activity in Golgi
membranes. Plant Physiol 130: 432–441
Ray PM (1980) Cooperative action of b-glucan synthetase and UDP-xylose
xylosyl transferase of Golgi membranes in the synthesis of xyloglucanlike polysaccharide. Biochim Biophys Acta 629: 431–444
Ray PM, Shininger TL, Ray MM (1969) Isolation of b-glucan synthetase
particles from plant cells and identification with Golgi membranes. Proc
Natl Acad Sci USA 64: 605–612
Reifenberger E, Freidel K, Ciriacy M (1995) Identification of novel HXT
genes in Saccharomyces cerevisiae reveals the impact of individual hexose
transporters on glycolytic flux. Mol Microbiol 16: 157–167
Richmond TA, Somerville CR (2000) The cellulose synthase superfamily.
Plant Physiol 124: 495–498
Robert S, Bichet A, Grandjean O, Kierzkowski D, Satiat-Jeunemaı̂tre B,
Pelletier S, Hauser MT, Höfte H, Vernhettes S (2005) An Arabidopsis
endo-1,4-b-D-glucanase involved in cellulose synthesis undergoes regulated intracellular cycling. Plant Cell 17: 3378–3389
Roberts AW, Roberts EM, Delmer DP (2002) Cellulose synthase (CesA) genes
in the green alga Mesotaenium caldariorum. Eukaryot Cell 1: 847–855
Saxena IM, Brown RM Jr (2005) Cellulose biosynthesis: current views and
evolving concepts. Ann Bot (Lond) 96: 9–21
Saxena IM, Brown RM Jr, Dandekar T (2001) Structure-function characterization of cellulose synthase: relationship to other glycosyltransferases. Phytochemistry 57: 1135–1148
Saxena IM, Brown RM Jr, Fevre M, Geremia RA, Henrissat B (1995)
Multidomain architecture of b-glycosyl transferases: implications for
mechanism of action. J Bacteriol 177: 1419–1424
Scheller HV, Ulvskov P (2010) Hemicelluloses. Annu Rev Plant Biol 61:
263–289
Sherson SM, Hemmann G, Wallace G, Forbes S, Germain V, Stadler R,
Bechtold N, Sauer N, Smith SM (2000) Monosaccharide/proton symporter AtSTP1 plays a major role in uptake and response of Arabidopsis
seeds and seedlings to sugars. Plant J 24: 849–857
Sims IM, Munro SLA, Currie G, Craik D, Bacic A (1996) Structural
characterisation of xyloglucan secreted by suspension-cultured cells of
Nicotiana plumbaginifolia. Carbohydr Res 293: 147–172
Somerville C (2006) Cellulose synthesis in higher plants. Annu Rev Cell
Dev Biol 22: 53–78
Sørensen I, Pettolino FA, Wilson SM, Doblin MS, Johansen B, Bacic A,
Willats WGT (2008) Mixed-linkage (1/3),(1/4)-b-D-glucan is not
unique to the Poales and is an abundant component of Equisetum
arvense cell walls. Plant J 54: 510–521
Szymanski DB, Cosgrove DJ (2009) Dynamic coordination of cytoskeletal
and cell wall systems during plant cell morphogenesis. Curr Biol 19:
R800–R811
Taylor NG (2008) Cellulose biosynthesis and deposition in higher plants.
New Phytol 178: 239–252
Taylor NG, Howells RM, Huttly AK, Vickers K, Turner SR (2003) Interactions among three distinct CesA proteins essential for cellulose
synthesis. Proc Natl Acad Sci USA 100: 1450–1455
Taylor NG, Laurie S, Turner SR (2000) Multiple cellulose synthase catalytic
subunits are required for cellulose synthesis in Arabidopsis. Plant Cell 12:
2529–2540
Timmers J, Vernhettes S, Desprez T, Vincken JP, Visser RGF, Trindade
LM (2009) Interactions between membrane-bound cellulose synthases
involved in the synthesis of the secondary cell wall. FEBS Lett 583:
978–982
Tiné MAS, Silva CO, de Lima DU, Carpita NC, Buckeridge MS (2006)
Fine structure of a mixed-oligomer storage xyloglucan from seeds of
Hymenaea courbaril. Carbohydr Polym 66: 444–454
Urbanowicz BR, Rayon C, Carpita NC (2004) Topology of the maize mixed
linkage (1/3),(1/4)-b-D-glucan synthase at the Golgi membrane. Plant
Physiol 134: 758–768
Vanzin GF, Madson M, Carpita NC, Raikhel NV, Keegstra K, Reiter WD
(2002) The mur2 mutant of Arabidopsis thaliana lacks fucosylated xyloglucan because of a lesion in fucosyltransferase AtFUT1. Proc Natl Acad
Sci USA 99: 3340–3345
Vergara CE, Carpita NC (2001) b-D-Glycan synthases and the CesA gene
family: lessons to be learned from the mixed-linkage (1/3),(1/4)b-Dglucan synthase. Plant Mol Biol 47: 145–160
Waldron KW, Brett CT (1983) A glucuronyltransferase involved in glucuronoxylan synthesis in pea (Pisum sativum) epicotyls. Biochem J 213:
115–122
Wang J, Elliott JE, Williamson RE (2008) Features of the primary wall
CESA complex in wild type and cellulose-deficient mutants of Arabidopsis thaliana. J Exp Bot 59: 2627–2637
Wang J, Howles PA, Cork AH, Birch RJ, Williamson RE (2006) Chimeric
proteins suggest that the catalytic and/or C-terminal domains give
CesA1 and CesA3 access to their specific sites in the cellulose synthase
of primary walls. Plant Physiol 142: 685–695
Weigel PH, DeAngelis PL (2007) Hyaluronan synthases: a decade-plus of
novel glycosyltransferases. J Biol Chem 282: 36777–36781
Wightman R, Turner SR (2008) The roles of the cytoskeleton during
cellulose deposition at the secondary cell wall. Plant J 54: 794–805
Williams KJ, Halkes KM, Kamerling JP, DeAngelis PL (2006) Critical
elements of oligosaccharide acceptor substrates for the Pasteurella
multocida hyaluronan synthase. J Biol Chem 281: 5391–5397
Wood PJ, Weisz J, Blackwell BA (1994) Structural studies of (1/3)(1/4)b-D-glucan by 13C-nuclear magnetic resonance spectroscopy and by
rapid analysis of cellulose-like regions using high-performance anion
exchange chromatography of oligosaccharides released by lichenase.
Cereal Chem 71: 301–307
Yong W, Link B, O’Malley R, Tewari J, Hunter CT, Lu CA, Li X, Bleecker
AB, Koch KE, McCann MC, et al (2005) Genomics of plant cell wall
biogenesis. Planta 221: 747–751
York WS, O’Neill MA (2008) Biochemical control of xylan biosynthesis:
which end is up? Curr Opin Plant Biol 11: 258–265
Yoshida M, Itano N, Yamada Y, Kimata K (2000) In vitro synthesis of
hyaluronan by a single protein derived from mouse HAS1 gene and
characterization of amino acid residues essential for the activity. J Biol
Chem 275: 497–506
Zabotina OA, van de Ven WTG, Freshour G, Drakakaki G, Cavalier D,
Mouille G, Hahn MG, Keegstra K, Raikhel NV (2008) Arabidopsis
XXT5 gene encodes a putative a-1,6-xylosyltransferase that is involved
in xyloglucan biosynthesis. Plant J 56: 101–115
Zeng W, Chatterjee M, Faik A (2008) UDP-xylose-stimulated glucuronyltransferase activity in wheat microsomal membranes: characterization
and role in glucurono(arabino)xylan biosynthesis. Plant Physiol 147:
78–91
Zeng W, Jiang N, Nadella R, Killen TL, Nadella V, Faik A (2010) A
glucurono(arabino)xylan synthase complex from wheat contains members of the GT43, GT47, and GT75 families and functions cooperatively.
Plant Physiol 154: 78–97
Zhong R, Morrison WH III, Freshour GD, Hahn MG, Ye ZH (2003)
Expression of a mutant form of cellulose synthase AtCesA7 causes
dominant negative effect on cellulose biosynthesis. Plant Physiol 132:
786–795
Zhong R, Peña MJ, Zhou GK, Nairn CJ, Wood-Jones A, Richardson EA,
Morrison WH III, Darvill AG, York WS, Ye ZH (2005) Arabidopsis fragile
fiber8, which encodes a putative glucuronyltransferase, is essential for
normal secondary wall synthesis. Plant Cell 17: 3390–3408
184
Plant Physiol. Vol. 155, 2011
Downloaded from on July 31, 2017 - Published by www.plantphysiol.org
Copyright © 2011 American Society of Plant Biologists. All rights reserved.