Running title: Synthesis of (1 β D-glycans* · 04/11/2010 · terminus of the acceptor glucan...

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Running title: Synthesis of (1→4)-β-D-glycans*

Title: Update on Mechanisms of Plant Cell Wall Biosynthesis: How plants make

cellulose and other (1→4)-β-D-glycans

Author: Nicholas C. Carpita

Department of Botany & Plant Pathology, and

Bindley Bioscience Center

Purdue University

915 West State Street

West Lafayette, IN 47907-2054

For correspondence:

Tel. +1-765-494-4653

FAX: +1-765-494-0363

E-mail: [email protected]

1This material is based upon work supported as part of the Center for Direct Catalytic

Conversion of Biomass to Biofuels (C3Bio), an Energy Frontier Research Center funded by the

U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, Award Number

DE-SC0000997

*Corresponding author; e-mail [email protected].

The author responsible for distribution of materials integral to the findings presented in this

article in accordance with the policy described in the Instructions for Authors

(www.plantphysiol.org) is: Nicholas C. Carpita ([email protected]).

Plant Physiology Preview. Published on November 4, 2010, as DOI:10.1104/pp.110.163360

Copyright 2010 by the American Society of Plant Biologists

https://plantphysiol.orgDownloaded on April 2, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

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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 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 for whose function is 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-synthase-like (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, to 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 differences

between plants with Type I walls, with those of the grasses with Type II walls (Fig. 1A; Penning

et al., 2009). For CesAs and certain Csl genes, establishment of specific function for the

synthases they encode comes from 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)-β-D- linkage, in which one



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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)-β-D- linkages.

Cellulose synthesis

In flowering plants, cellulose is a para-crystalline array of about two to three dozen (1→4)-β-D-

glucan chains. Microfibrils of 36 glucan chains have a theoretical diameter of 3.8 nm, but X-ray

scattering and NMR spectroscopy indicate 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 particle 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. 2B 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)-β-D-glycans (Saxena et al., 1995). The

higher-plant CesA genes are predicted to encode polypeptides of about 110 kDa, 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).



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Further evidence for the function of plant CesA genes in cellulose synthesis came from

Arabidopsis mutants of three of the CesA genes involved in primary wall synthesis: a

temperature-sensitive radial swelling mutant rsw1 (AtCesA1; Arioli et al., 1998), a dwarf-

hypocotyl mutant procuste (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, 2002; Caño-Delgado et al., 2003). Despite co-expression 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., 2001, 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 non-redundant 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 knock-outs of AtCesA3 (Ellis and Turner, 2002; Caño-Delgado et al., 2003) and AtCes6

(Fagard et al., 2000) result in partially impaired synthesis but not total inhibition. Desprez et al.

(2007) indicate 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 co-

localization 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-terminus 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 homo-

and heterodimers of AtCesA1, AtCesA3 and AtCesA6 are inferred. Wang et al. (2008) used pull-



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down experiments similar to those of Taylor et al. (2003) to show that these three primary wall

CesA proteins interact. Further, they showed that Triton-soluble microsomal preparations

subjected to native PAGE gave an 840 kDa complex, and that null mutants, but not missense

mutations, gave smaller 420 kDa complexes (Wang et al., 2008). Atanassov et al. (2009)

affinity-trapped a ladder of complexes of CesA oligomers to about 700-730 kDa. 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)-β-D-glucan chain require one or two catalytic polypeptides?

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; Guerreiro et al.,

2010), with only a few reports of cellulose synthesis in vitro with isolated membranes (e.g.

Kudlicka and Brown, 1997; Lai-Kee-Him et al., 2002). For both cellulose and the related (1→4)-

β-D-glycan, chitin, synthesis proceeds by attachment of glucosyl residues to the non-reducing

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)-β-D- linkage means that each glucosyl residue is turned 180° with respect to

each neighbor. Thus, the O-4 position of non-reducing terminal sugar of the acceptor chain is

displaced several Ångstroms upon addition of each successive unit (Fig. 3B). For the next

glycosyl transfer to occur, then the site of catalysis must move several Ångstroms 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 (Vergara and Carpita, 2001;

Buckeridge et al., 2001). In either model, glycobiosyl units, or any even-numbered oligomeric

units, are added to the non-reducing end to ensure that the (1→4)-β-D- linkages are strictly



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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 polysaccharide synthases indicate that a single polypeptide is sufficient. Hyaluronan (HA)

is an unbranched polysaccharide composed of repeating units of (1→3)-β-D-N-

acetylglucosamine and (1→4)-β-D-glucuronic acid (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 synthesis of a single HA polymer

requires only a single polypeptide.

The crystal structure of a non-processive 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.

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 synthesis of a repeating

(1→4)-β-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 as 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 from the crystal structure of the SpsA synthase (Charnock and Davies,

1999). This is the first direct evidence by crystal structure of a homodimer formed by GT2



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proteins, but even more complicated structures are also found. For example, an E. 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 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, Purdue University, 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 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 have both activities on a

single peptide does not preclude the possibility that 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 8 membrane spans

proposed for the cellulose synthase of sufficient size to extrude a (1→4)-β-D-glucan chain. The

question about sufficient channel size was raised also with respect to the GT family 2 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



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(Pao et al., 1998; Reifenberger et al., 1995; Sherson et al., 2000; Klepek et al., 2009).

The sensitivity of detergent-solubilized CesAs complexes to DTT suggested to Atanassov et

al. (2009) that disulfide bonds are involved in the coupling into the 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 Zn 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

thioredoxin-like 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 β-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 H2O2, suggesting 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 HVR 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 distinct sub-clade structure. Potential protein-

protein interactions through sub-domains of this region containing conserved Cys residues,

clusters of consecutive basic Lys and Arg residues, and clusters of acidic Asp and Glu residues

forming the basis of a Class Specific Region (Fig. 1B; Vergara and Carpita, 2001).



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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, S. Ghosh, unpublished results). 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 homo- or hetero-dimers 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 non-reducing 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)-β-D-glucan chains instead of 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 thirty minutes, 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 suspected to be necessary for the formation

of primers of polymer synthesis, metabolic channeling of substrates, chain crystallization of the

chains, and termination of chains (Doblin et al., 2002; Somerville, 2006; Guerreiro et al., 2010).

Further, 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 mimick (Glu) phoshorylation have multiple effects that either reduce synthesis rates or

interactions with the microtubule cytoskeleton independently (Chen et al., 2010).



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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 sucrose 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. β-Glucan microfibrils are synthesized from sucrose and UDP

on immobilized tobacco 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 glucose 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 sitosterol-cellodextrins synthesized from

sitosterol-β-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 find double mutants of two major sterol-β-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 the KORRIGAN

(Robert et al., 2005) might represent a dynamic factory associated with both the microtubule

network and 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, 2006; Gutierrez et al., 2009). These studies also established



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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

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 Golgi-derived vesicles to the sites of cellulose synthesis much progress has already been made

along these lines (Held et al., 2008; Konopka and Bednarek, 2008; Crowell et al., 2009).

The synthesis of non-cellulosic polysaccharides with (1→4)-β-D-glycan backbones

Because of the same conserved domains of nucleotide-sugar binding and catalysis as those

encoding CesAs, sub-families of Csl genes were predicted to encode the synthases of non-

cellulosic polymers with (1→4)-β-D-glycan backbones, primarily (gluco)mannans and

galacto(gluco)mannans, xyloglucans, glucuronoarabinoxylans (GAX), and the grass-specific

(1→3),(1→4)-β-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 β-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)-β-D-glucan synthase is the topological

equivalent of cellulose synthase

The mixed-linkage (1→3),(1→4)-β-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

they probably arose by convergent evolution of synthases. For the grasses, this glucan is not a

random mixture of (1→3)-β-D- and (1→4)-β-D-glucosyl linkages, but is composed primarily of

cellotriosyl and cellotetraosyl units in a ratio of about 2.5:1 connected by single (1→3)-β-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



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abundance than the next even-numbered unit in the series. The synthesis of the (1→3),(1→4)-β-

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

μM concentrations of UDP-[14C]-glucose 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 μM (Buckeridge et al., 1999), indicating that the mechanism of synthesis of the odd-

numbered cellodextrin unit is fundamentally different than 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)-β-

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 glycosyl transferase (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)-β-D-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)-β-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 odd-numbered 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)-β-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)-β-D-

glucan synthase (Buckeridge et al., 2001) ‘default’ to synthesis of the (1→3)-β-D-glucan,



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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 (Nishamura et al., 2003), only Golgi membranes from grasses–

the only angiosperms that make the (1→3),(1→4)-β-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)-β-D-glucan in vitro as well as (1→4)-β-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)-β-D-glucan biosynthesis. Heterologous

expression of a rice CslF in Arabidopsis, a species that does not make (1→3),(1→4)-β-D-glucan,

results in small amounts of the β-D-glucan in the cell walls (Burton et al., 2006). However,

considerably greater amounts of the (1→3),(1→4)-β-D-glucan result when a CslH is co-

expressed 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 cross-linking glycans might be

encoded by Csls 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)-β-D- linkages of cellulose, and UDP-Glc is

the substrate for (1→3)-β-D-glucans (Chambers and Elbein, 1970), had already been disproven.



14

Still today GDP-Glc is still listed erroneously as the substrate for cellulose synthesis on most

wall charts of biochemical pathways, even though 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 stimulation of incorporation into a glucomannan product (Elbein

and Hassid, 1966; Piro et al., 1993).

The knowledge of GDP-nucleotide sugars as substrates were instrumental in the discovery

that a CslA gene encodes a mannan synthase by expression profiling of guar 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 sub-groups 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 Arabidopsis-dominated and the

mixed sub-group (Penning et al., 2009). The functions of these other sub-group 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, 1980). Small amounts of xyloglucan-like

oligomers with the characteristic α-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, 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 is consistent with an even-number cellobiose



15

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 4-5-5-4 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 UDP-Glc for backbone

synthesis of (1→4)-β-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)

propose 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. (2006) provide evidence that

CslC genes encode the synthases of the xyloglucan backbone. Although Pichia is unable to

make UDP-Xyl, co-expression of the xylosyl transferase with CslC is sufficient to induce

extension of (1→4)-β-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 sub-family 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., 2009)

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 α-L-Fuc-

(1→2)-β-D-Gal-(1→2)-α-D-Xyl- trisaccharide extension (Bauer et al., 1973), but others, such as



16

those of Solanaceous species, have a truncated unit structure substituted with α-L-Ara-(1→2)-α-

D-Xyl 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). Further, the

xyloglucans synthesized in the Golgi are modified in ways that make them structurally different

than those that are assembled onto the cellulose microfibrils in the wall (Obel et al., 2009).

The purification of a xyloglucan-specific fucosyl transferase that led to the discovery of a

glycosyl transferase family 37 (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 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)-β-D-xylans from UDP-Xyl 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)-β-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; Porchia et al., 2002; Kuroyama and Tsumuraya, 2001; Zeng et

al., 2008).

Apart from a hint that the AtCslD5 may be involved (Bernal et al., 2007) in the synthesis of

(1→4)-β-D-xylans, 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)-β-D-Glucuronoxylan (GX) synthesis appears to involve a complex initiation

sequence. Peña et al. (2007) discovered that collapsed xylem mutants deficient in xylan, irx8



17

and fra8, were essentially devoid of a complex tetrasaccharide, β-D-Xyl-(1→3)-α-L-Rha-(1→2)-

α-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

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)-β-D-xylan 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) encode members of the GT8 of Group C, and

IRX7/FRA8 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)-β-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 to the anomeric linkage in the nucleotide sugar, so GT8 members make α-D-linkages,

whereas the GT47 members make β-D- or α-L- linkages. While an α-D-GalA is found in the

tetrasaccharide, (1→2)-α-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 addition of these GlcA side-groups play 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 been ruled out. In their model a type of alternating

‘pendulum’ mechanism is proposed for the introduction of the (1→4)-β-D-xylan linkages, a

mechanism analogous to the dimer synthesis described here for (1→4)-β-D- linkages in general.



18

We are just learning the identity of the GlcA and Araf transferases of the GX and GAX polymers

and the distinctions between the GX devoid of arabinose 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 on 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

preservation of synthesis (Carpita and Delmer, 1980). This work was extended to show that

artificial potentials stimulate (1→3)-β-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)-β-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



19

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 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).



20

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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 non-reducing

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-groups. 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-

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