Download Biochemistry and physiological roles of enzymes that `cut and paste

Journal of Experimental Botany, Vol. 64, No. 12, pp. 3519–3550, 2013
doi:10.1093/jxb/ert201 10.1093/jxb/ert201
Darwin Review
Biochemistry and physiological roles of enzymes that ‘cut
and paste’ plant cell-wall polysaccharides
Lenka Franková and Stephen C. Fry*
The Edinburgh Cell Wall Group, Institute of Molecular Plant Sciences, School of Biological Sciences, The University of Edinburgh, The
King’s Buildings, Mayfield Road, Edinburgh EH9 3JH, UK
* To whom correspondence should be addressed. E-mail: [email protected]
Received 15 April 2013; Revised 30 May 2013; Accepted 4 June 2013
Abstract
The plant cell-wall matrix is equipped with more than 20 glycosylhydrolase activities, including both glycosidases
and glycanases (exo- and endo-hydrolases, respectively), which between them are in principle capable of hydrolysing
most of the major glycosidic bonds in wall polysaccharides. Some of these enzymes also participate in the ‘cutting
and pasting’ (transglycosylation) of sugar residues—enzyme activities known as transglycosidases and transglycanases. Their action and biological functions differ from those of the UDP-dependent glycosyltransferases (polysaccharide synthases) that catalyse irreversible glycosyl transfer. Based on the nature of the substrates, two types of
reaction can be distinguished: homo-transglycosylation (occurring between chemically similar polymers) and heterotransglycosylation (between chemically different polymers). This review focuses on plant cell-wall-localized glycosylhydrolases and the transglycosylase activities exhibited by some of these enzymes and considers the physiological
need for wall polysaccharide modification in vivo. It describes the mechanism of transglycosylase action and the
classification and phylogenetic variation of the enzymes. It discusses the modulation of their expression in plants at
the transcriptional and translational levels, and methods for their detection. It also critically evaluates the evidence
that the enzyme proteins under consideration exhibit their predicted activity in vitro and their predicted action in vivo.
Finally, this review suggests that wall-localized glycosylhydrolases with transglycosidase and transglycanase abilities
are widespread in plants and play important roles in the mechanism and control of plant cell expansion, differentiation, maturation, and wall repair.
Key words: Cell expansion, cell wall, glycanases, glycosidases, hemicelluloses, hydrolysis, oligosaccharides, pectins,
remodelling, transglycanases, transglycosidases, transglycosylation, xyloglucan.
Introduction
A primary wall layer is (or was, at the time of its formation)
susceptible to plastic extension: i.e. able to accommodate the
cell’s irreversible expansion. Secondary wall layers, in contrast, which may be deposited internal to the primary wall
after cell expansion has ceased and have quite distinct chemical compositions, do not subsequently increase in area. The
primary wall thus serves the key role of defining the shape
and size of the plant cell.
Plant primary cell walls are not rigid or inert ‘boxes’ but
constitute a flexible and metabolically active extraprotoplasmic compartment; they control cell expansion by varying their
extensibility. This ability to change biophysically is conferred
by biochemical reactions and molecular rearrangements that
occur within the walls (in muro).
The constituent polysaccharides of plant cell walls are
synthesized by the protoplast, mostly within Golgi bodies
except that cellulose and callose are produced at the plasma
membrane. After synthesis in the Golgi body, polysaccharides are carried in vesicles to the plasma membrane to
be deposited by exocytosis on the inner face of the existing wall. In muro, the polysaccharides may undergo many
interesting and physiologically relevant modifications,
© The Author [2013]. Published by Oxford University Press on behalf of the Society for Experimental Biology.
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3520 | Franková and Fry
including transglycosylation (‘cutting and pasting’ molecules), cross-linking, and hydrolysis. Other proposed wallremodelling reactions include transacylation (of polymers
possessing carboxy groups) and non-enzymic scission
caused by hydroxyl radicals. By means of this repertoire of
polysaccharide modifications, the cell has the subtlety to
manipulate the ‘nuts and bolts’ of development, especially
the direction and rate of cell growth, by remodelling wall
polysaccharides.
Physiological roles of the primary cell wall that can be
finetuned by the in-muro metabolic reactions discussed
in this review include not only dictating cell shape and
size but also causing programmed leaf abscission, pod
dehiscence for seed dispersal, seed-coat bursting during
germination, becoming ‘perforated’ in the case of xylem
and phloem, supplying food reserves for seedling growth,
generating and subsequently inactivating oligosaccharin signals, governing the wall’s porosity and ability to
adsorb metal ions, and defending the cell against microbial ingress.
Chemical composition of the primary
cell wall
To understand wall remodelling, we need a summary of primary cell-wall chemistry. In land plants, polysaccharides constitute the bulk of this wall’s dry mass, grouped into three
broad classes: cellulose, hemicelluloses, and pectins (Scheller
and Ulvskov, 2010; Albersheim et al., 2011; Fry, 2011). In
dicot primary walls, these three classes occur at very roughly
1:1:1 by weight. Cellulose forms partially crystalline microfibrils, the wall’s skeleton, devoid of internal water; hemicelluloses hydrogen-bond strongly to the cellulose and may also
become locally trapped within microfibrils; and pectins plus
hemicelluloses together constitute a hydrated matrix occupying the space between microfibrils (Fig. 1). In a widely adopted
(but speculative) model, long rope-like hemicellulose chains
adhere to multiple microfibrils and tether them, restraining
cell expansion (Fry, 1989; Hayashi, 1989). Recent critical
evaluation (Park and Cosgrove, 2012) suggests that, although
correct in outline, this model is an oversimplification.
Fig 1. Model of the polysaccharide framework in a plant cell wall, generalized for poalean and non-poalean walls. 1, Cellulose:
cellulose microfibrils; 2–6, hemicelluloses: 2, xyloglucan; 3, mixed-linkage glucan; 4, xylan and related heteroxylans; 5, callose; 6,
mannan and related heteromannans; 7–11, Pectins: 7, galactan; 8, arabinan; 9, homogalacturonan; 10, rhamnogalacturonan I; 11,
rhamnogalacturonan II; 12, boron bridge; 13, ‘egg-box’ with calcium bridges; 14–16, Non-polysaccharide components: 14, enzymes
and structural proteins; 15, cellulose synthase complex; 16, transport vesicles.
Plant cell-wall enzymes | 3521
Cellulose is a chain of (1→4)-linked β-Glc residues lacking
side chains.1 An elementary microfibril (produced by a single
‘rosette’ of cellulose synthases) consists of ~16–18 cellulose
molecules lying in parallel (Guerriero et al., 2010). Cellulose
cannot be extracted from the wall into aqueous solution
except by aggressive complexing agents such as cadoxen (10%
cadmium oxide in aqueous 30% 1,2-diaminoethane).
Hemicelluloses are polymers with a backbone of β-Glc,
β-Xyl, or β-Man (or β-Man and β-Glc), mainly or entirely
(1→4)-linked (except callose), and usually possessing short
side chains; most are neutral. Highly significant hemicelluloses
of the primary walls of all vascular plants are xyloglucans
(backbone β-(1→4)-Glc; side chains mainly α-Xyl, and often
also β-Gal-(1→2)-α-Xyl and α-Fuc-(1→2)-β-Gal-(1→2)-αXyl, attached at position 6 of some of the Glc residues) and
xylans sensu lato (backbone β-(1→4)-Xyl; side chains usually
α-Araf and/or α-GlcA attached at positions 2 and/or 3 of some
of the Xyl residues (the polysaccharides may then be more precisely referred to as arabinoxylans, glucuronoarabinoxylans,
etc., some being acidic). Xyloglucans predominate in noncommelinid species; xylans in commelinids. Sugar sequences
within xyloglucan chains can be specified by a series of code
letters, comparable to those used in reporting sequences of
proteins or DNA; for details see Fry et al. (1993). For example, G = β-Glc of the backbone, not further substituted;
X = α-Xyl-β-Glc; L = β-Gal-(1→2)-α-Xyl-β-Glc; F = α-Fuc(1→2)-β-Gal-(1→2)-α-Xyl-β-Glc. All 18 xyloglucan code-letters currently in use are listed by Franková and Fry (2012b).
Frequently observed sequences in dicot xyloglucans include
XXXG, XXLG, XXFG, and XLFG. A comparable system of
abbreviations is in use for xylan sequences (Fauré et al., 2009).
Relatively minor primary wall hemicelluloses in most land
plants, but predominant in eusporangiate ferns, are mannans
sensu lato (backbone β-(1→4)-Man, often with interspersed
β-(1→4)-Glc; sidechains often α-Gal on position 6 of some
of the Man residues). An additional vascular land-plant
hemicellulose, restricted to but abundant in commelinids
(grasses, cereals, etc.) and Equisetum (horsetails) (see also
the section ‘Phylogenetic variation’), is mixed-linkage glucan
(MLG) (backbone β-(1→4)-Glc interspersed with a minority
of β-(1→3)-Glc residues; no side chains).
Although chemically diverse, hemicelluloses generally share
an ability to hydrogen-bond to cellulose and are extractable
from the cell wall in alkali (optimally 6 M NaOH at 37 °C;
Edelmann and Fry, 1992).
Pectins (Fig. 1) are α-GalA-rich polysaccharides built up of
distinct domains, the simplest of which is homogalacturonan,
a linear homopolymer of (1→4)-linked α-GalA residues, partially methyl-esterified and often also partially O-acetylated.
More complex pectic domains are the rhamnogalacturonans
(RG-I and RG-II) and xylogalacturonan. It is widely thought
that these pectic domains are glycosidically bonded, end-to-end,
into a complete polysaccharide, conveniently described simply
as ‘pectin’. Details of pectin structure are given by Albersheim
et al. (2011) and Fry (2011). In brief, RG-I has a backbone of
the repeating disaccharide, ...(1→4)-GalA-(1→2)-Rha-..., with
some of the Rha residues carrying neutral oligosaccharide side
chains rich in Gal and/or Araf. RG-II is a small domain with a
backbone of (1→4)-linked α-GalA residues with highly complex
oligosaccharides linked to the backbone via Apif residues; in the
cell wall, RG-II domains are usually cross-linked to each other
via borate diester groups attached to one of the Api residues.
Primary wall polysaccharides may possess, in addition to
sugar residues, non-carbohydrate substituents: for example,
acetyl, feruloyl, and methyl esters. Other non-carbohydrate
components of certain primary walls include (glyco)proteins,
lignin, cutin, suberin, and silica. It is often incorrectly stated
that primary walls do not lignify; however, lignification of
xylem vessels begins in their primary walls.
The ratio of the various primary wall polysaccharides differs between taxa and between tissues, and varies developmentally within a given cell. For example, during maturation,
MLG increases in Equisetum (Fry et al., 2008b) but decreases
in commelinids (Buckeridge et al., 2004). Pectins are major
components of all fast-growing cells except in commelinids,
which are particularly rich in xylans (Carpita and Gibeaut,
1993). Mannans are highly abundant in some fern allies and
are also present in many algal species, some of which completely lack cellulose (Popper and Fry, 2004). Several other
important shifts in wall polysaccharide chemistry accompanied major events in plant evolution. For example, xyloglucans are present in all land plants examined but have not been
chemically demonstrated in any algae – including the charophytes,2 from which all land plants originated – implying that
the ‘invention’ of xyloglucan accompanied (and may have
enabled) the invasion by plants of the land. Wall-remodelling
enzymes targeting the major wall polysaccharides thus need
to be tailored to taxa, tissues, and developmental stages.
The physiological need for wall
polysaccharide modification in vivo
Cell expansion: hydrolysis of wall components
The primary wall confers the cell’s ability to define its own shape
and size. Cell expansion, an irreversible increase in cell volume
often exceeding 1000-fold, is a dramatic and unique feature
of plants, with no equivalent in animals or most micro-organisms. Controlled plant cell expansion demands the reversible
‘loosening’ of the cellulose–hemicellulose–pectin primary
1
Sugar abbreviations: Api, d-apiose; Ara, l-arabinose; Fuc, l-fucose; Gal, d-galactose; GalA, d-galacturonic acid; Glc, d-glucose; GlcA, d-glucuronic
acid; Man, d-mannose; Rha, l-rhamnose; Xyl, d-xylose. All sugars are in the six-membered pyranose ring form unless marked ‘f’ for furanose.
2
The status of xyloglucan in charophytes is controversial. Immunological evidence supports the presence of a xyloglucan-like polymer in Chara,
Coleochaete, Cosmarium, and Netrium (Van Sandt et al., 2007; Sørensen et al., 2011), but enzymic digestion of Chara and Coleochaete cell walls has
consistently failed to yield xyloglucan’s diagnostic disaccharide, isoprimeverose (e.g. Popper and Fry, 2003). Methylation analysis of Spirogyra cell walls
demonstrated 4-linked and 4,6-linked Glc and terminal Xyl residues (Ikegaya et al., 2008), but these residues’ anomerism was not determined and the
Glc residues could have been α-Glc of starch (which is often difficult to completely remove by amylase digestion) rather than β-Glc of xyloglucan, and
the terminal Xyl residues could have been β-Xyl from xylans rather than α-Xyl of xyloglucan. Resolution of these discrepancies requires further research.
3522 | Franková and Fry
wall. Thus, plants will require a battery of wall-manipulating
enzymes not found in other organisms (Labavitch, 1981; Fry,
1995, 2004; de la Torre et al., 2002; Minic, 2008). Besides
the intrinsic interest of these wall enzymes for understanding plant growth, they are also exciting subjects for genetic
manipulation and targets for novel herbicides. Enzymes that
loosen the primary wall, for example by catalysing its partial
hydrolysis, may be expected to step up the growth rate.
Wall assembly: recruiting new polysaccharides
Roles of polysaccharide-remodelling enzymes are not confined to cell expansion. For example, enzyme activities that
‘cut and paste’ glycosidic bonds (transglycanases, e.g. xyloglucan endotransglucosylase, XET) can create new polysaccharide–polysaccharide linkages and thus play a role in recruiting
newly secreted polysaccharides into the wall fabric, contributing to wall assembly. In addition, hetero-transglycanases can
graft part of one polysaccharide to a qualitatively different
one, which may also contribute to wall assembly.
Wall loosening and/or strengthening: rewiring
glycosidic bonds in old polysaccharide chains
The cutting/pasting of polysaccharide chains by transglycanases
occurs not only at the moment of secretion but also between
pairs of polysaccharides which have already been part of the
wall architecture for some time. It is difficult to deduce whether
such reactions contribute predominantly to wall loosening and
thus growth promotion (Thompson and Fry, 2001) or to wall
strengthening by stitching polymers together, (e.g. in xylem
cell walls; Nishikubo et al., 2007). Other homo-transglycanase
reactions include the cutting/pasting of xylans to xylans and of
mannans to mannans; precise physiological roles for these processes are unclear, but may include reserve mobilization after
germination of mannan-rich seeds and wall loosening during
growth and fruit softening. In addition, a hetero-transglycanase
activity, MLG:xyloglucan endotransglucosylase (MXE), can
link a portion of MLG to a xyloglucan chain, thus creating a
chimaeric polymer with MLG at one end and xyloglucan at the
other. This may contribute to strengthening the Equisetum stem
in mature plants, helping it to resist wind damage or herbivory.
Abscission/dehiscence
Another role for wall polysaccharide modification concerns
enzymes that disrupt tissue cohesion by lysing the primary
wall and/or middle lamella, thus permitting cell–cell separation (Le Cam et al., 1994; González-Carranza et al., 2007;
Zhang et al., 2007). This enables: leaf abscission in deciduous
trees; dehiscence of pod-like fruits, e.g. follicles, legumes, siliquae, and capsules; softening in drupes and berries; and the
spatially and temporally targeted rupture of the endosperm
and/or seed coat at the moment of germination.
Perforations
On a highly localized scale, plant enzymes can bore holes
in cell walls, for example during the formation of xylem
perforation plates, phloem sieve plates, and plasmodesmata,
enabling the intercellular transport of xylem sap, sucrose,
and RNAs, respectively (Bollhöner et al., 2012; Tilsner and
Oparka, 2012).
Wall polysaccharide mobilization
Enzymic lysis also occurs in the walls of certain seeds, whose
enormous stockpiles of specific polysaccharides such as
xyloglucans or galactomannans are hydrolysed, ultimately
to monosaccharides, providing a carbon and energy source
for the seedling until it attains photosynthetic self-sufficiency.
Related to seed reserves, it would a priori appear energy-efficient for deciduous trees to degrade some leaf polysaccharides
shortly before abscission and to salvage the resultant sugars
by basipetally transporting them via the phloem (Hoch,
2007); more work is required, however, to test this idea.
Making oligosaccharins
On a much smaller scale, but nevertheless qualitatively significant, some wall polysaccharides undergo enzymic turnover
to release biologically active oligosaccharides (‘oligosaccharins’) with putative signalling roles (McDougall and Fry,
1991; Darvill et al., 1992; Aldington and Fry, 1993; BeňováKákošová et al., 2006). For example, fragments of xyloglucan possessing an α-Fuc residue can at concentrations of
1 nM antagonize the cell expansion that is induced by 1 μM
auxin. Conversely, higher concentrations (0.1–10 μM) of
several xyloglucan oligosaccharides can promote cell expansion, and such concentrations are indeed present in the apoplast in vivo, at least in cell-suspension cultures (Fry, 1986).
Glucomannan-related oligosaccharins have been reported to
promote cell elongation in roots but inhibit it in hypocotyls
(Richterová-Kučerová et al., 2012). Other oligosaccharins,
produced at least in vitro from homogalacturonan, antagonize auxin-induced growth (Ferrari et al., 2008) and trigger
wound-hormone release; however, such oligogalacturonides
appear more likely to be generated by the action of microbial
enzymes than by the plant’s own repertoire.
Removing oligosaccharins
In addition to enzymes generating oligosaccharins from
polysaccharides, plants also possess enzymes that degrade
oligosaccharins, either by hydrolysis or by grafting large poly­
sa­ccharides to them (Baydoun and Fry, 1989; Darvill et al.,
1992; García-Romera and Fry, 1995). This may be important
since biological ‘messages’ need to be inactivated when the
information that they carry is no longer relevant to the plant’s
environmental or developmental situation.
Wall porosity
Partial degradation of certain cell-wall polysaccharides, especially pectins, can finetune the molecular pore size of the wall
fabric (Carpita et al., 1979; Baron-Epel et al., 1988), thereby
modulating what sizes of molecules, such as arabinogalactanproteins, can potentially move intercellularly as signals.
Plant cell-wall enzymes | 3523
Cell defence
Glycosidase and glycanase activities
Finally, the wall contains enzymes that may render it impenetrable by potential pathogens or indigestible by herbivores.
Such enzymes would be mainly those involved in cross-linking,
for example peroxidases and phenol oxidases (laccases). To the
list of wall-bolstering wall enzymes could be added the newly
discovered acyltransferase, cutin synthase (Yeats et al., 2012).
An enzyme catalysing the hydrolysis of poly- and/or oligosaccharides will attack either at terminal sugar residues (almost
always non-reducing termini) or at mid-chain residues, not
both. Such exo- and endo-glycosyl hydrolase activities are
termed ‘glycosidases’ and ‘glycanases’, respectively. Note
also the distinction between ‘glyc-’, referring to an unspecified sugar residue and ‘gluc-’, referring to a glucose residue.
Examples of glycosidases are α-xylosidase, β-xylosidase,
and β-glucosidase; examples of glycanases include (1→4)β-xylanase and (1→4)-β-glucanase (cellulase). In the following sugar sequences, the reducing terminus is on the right.
An enzyme that splits off the non-reducing terminal β-Xyl residue from the model substrate
(1→4)-β-xyloh­exaose (Xyl6) is a β-xylosidase (Fig. 2A):
Glycosidic bonds and enzyme activities
that act on them
In this work, the term ‘enzyme’ follows the Enzyme Commission
usage: it is defined as an ‘activity’, which, however, may be
shared by multiple isozymes encoded by different genes.
Fig. 2. Diagrammatic representations of the activities of glycosidases (A, B), glycanases (C), transglycosidases (D, E), and
transglycanases (F), Each circle represents a sugar residue; , bond cleaved.
3524 | Franková and Fry
We will show the reactants and products in the order:
donor + acceptor ↔ hybrid product + leaving group.
It is helpful to distinguish readily reversible transglycosylation reactions, in which the bond cleaved has an energy (ΔG0′;
i.e. free energy of hydrolysis) similar to that of the bond
formed, from essentially irreversible ones (as will be discussed
in the next section). Reversible transglycosylation is appropriate for wall remodelling, in which the reaction does not
proceed in any defined direction.
By analogy with hydrolase names, we use the terms ‘transglycosidase’ and ‘transglycanase’ for exo- and endo-enzymes
respectively, referring to the glycosidic bond that is cleaved.
Thus, the reactions in Fig. 2D–F represent trans-β-xylosidase,
trans-β-galactosidase, and trans-β-xylanase, respectively (an
alternative nomenclature uses ‘xylan endotransglycosylase’
instead of trans-β-xylanase; Johnston et al., 2013).
When the donor is qualitatively similar to the acceptor, the
reaction is ‘homo-transglycosylation’. Examples include the
activities of trans-β-mannanase, trans-β-xylanase, and xyloglucan endotransglucosylase (XET,3 ‘trans-xyloglucanase’;
Fig 4). When the donor and acceptor are qualitatively different (hetero-transglycosylation), the nomenclature is more
complex since both the donor and the acceptor have to be
specified. A convenient system for naming such activities
takes the form ‘donor:acceptor endotransglycosylase’, e.g.
MLG:xyloglucan endotransglucosylase (MXE; Fig 4).
Xyl-Xyl-Xyl-Xyl-Xyl-Xyl + H2O → Xyl + Xyl-Xyl-Xyl-Xyl-Xyl.
An enzyme that cleaves a mid-chain linkage in the same
substrate is a β-xylanase (Fig. 2C), for example:
Xyl-Xyl-Xyl-Xyl-Xyl-Xyl + H2O → Xyl-Xyl-Xyl-Xyl + Xyl-Xyl.
The terms ‘glycosidase’ and ‘exo’ do not imply that the
attacked terminal residue is necessarily located at the end of
the backbone; side chains are also non-reducing termini, for
example certain β-Gal residues of xyloglucan attacked by
β-galactosidase (Fig. 2B). Glycanases catalyse the hydrolysis
of glycosidic bonds within the backbone of the polysaccharide (Fig. 2C) or mid-chain bonds within a lengthy side chain.
Glycosidases generally have high specificity for the glycosyl
group attacked, but there is often a lower specificity towards
the nature of the ‘aglycone’ (for explanation, see Fig. 3A).
For example, β-xylosidase may release monomeric xylose
from the non-reducing terminus of xylan (Xyln), xylohexaose
(Xyl6; Fig. 3c), and even p-nitrophenyl β-xyloside (Fig. 3B).
However, there are important cases of tighter specificity: for
example, plant α-xylosidases release xylose from the 1st but
not the 2nd or 3rd (counting from the non-reducing end) Xyl
residue of the xyloglucan heptasaccharide XXXG and do not
attack p-nitrophenyl α-xyloside; Fanutti et al., 1991).
Transglycosidase and transglycanase activities: readily
reversible
Transglycosylation is a ‘cutting and pasting’ reaction in which
a glycosidic bond is cleaved, but not by hydrolysis. Instead, the
broken bond’s energy is conserved in forming a new glycosidic
linkage (Fig. 2D–F). The substrates are a donor and an acceptor; the products are a ‘hybrid’ and a leaving group (Fig. 2D).
‘Irreversible’ transglycosylation
Essentially irreversible transglycosylation reactions make the
glycosidic bonds during de-novo polysaccharide synthesis.
Aglycone
(oligosaccharide)
a
b
Aglycone
(p-nitrophenol)
Aglycone
(general)
H
H
HO
H
OH
H
O
H
OH
O
H
H
H
HO
H
OH
H
O
H
OH
c
NO2
H
H
Glycosidic
oxygen
R
O
H
H
H
H
H
HO
H
OH
H
O
O
H
OH
H
OH
H
H
O
O
H
OH
H
H
OH
H
O
H
H
OH
OH
n
Glycosidic bond
to be cleaved
Anomeric centre
Glycosyl group
(β-Xylosyl)
Glycosyl group
(β -Xylosyl)
Glycosyl group
(β -Xylosyl)
Fig. 3. Glycosyl groups versus aglycones, and the bonds cleaved by glycosidases and transglycosidases. In each case, the aglycone is
released (with an H atom in place of the red bond). Green arrow, glycosidic oxygen atom; red line, glycosidic bond to be cleaved by the
glycosidase; black arrow, anomeric centre of the glycosyl group under consideration. Note that an aglycone can itself be a sugar.
3
‘XET’ was defined by Rose et al. (2002) as ‘xyloglucan endotransglucosylase’ activity, which is a more precise and informative term that the
previously used ‘xyloglucan endotransglycosylase’. Since then, ‘endotransglucosylase’ has been criticized (Eklöf and Brumer, 2010) on the
grounds that the enzyme transfers a whole length of the polysaccharide chain, not just a single Glc residue. However, this is not a valid criticism
because the distinction between ‘gluco’ and ‘glyco’ does not concern the number of residues transferred but their identity. ‘Transglucosylase’
specifies that a Glc–X bond is cleaved and reformed; a ‘transglycosylase’ could act on a Xyl–X or Gal–X bond, and is thus a term that might be
used when the nature of the cleaved/reformed bond is uncertain.
Fig. 4. Proposed biological roles of enzymes with transglycosidase and transglycanase activities that catalyse transglycosylation reactions between oligosaccharides and/or
polysaccharides of the plant cell wall.
Plant cell-wall enzymes | 3525
3526 | Franková and Fry
Here, the bond broken is more ‘energy-rich’ (has a larger negative ΔG0′ of hydrolysis) than the one made. Irreversibility
is appropriate for biosynthesis of major metabolic end products such as wall polysaccharides. The high-energy donors
are usually NDP-sugars (e.g. UDP-xylose), the acceptor is
the nascent polysaccharide, and the enzymes are polysaccharide synthases or ‘NDP-sugar : glycan glycosyltransferases’.
This type of enzyme is membrane-bound and not discussed
in detail in this review.
Carbohydrate-Active enZyme proteins and
their genes
Browsing for plant ‘cutting-and-pasting’ enzymes
We will now describe specific examples of potential wallremodelling enzymes, focusing on transglycosidases and transglycanases. Thanks mainly to recent progress in proteomics,
genomics, and metabolomics, numerous enzyme databases
have been created and are now available online. However,
this enormous resource is significantly reduced when one is
searching for plant enzyme databases, especially when focusing on those that provide information on carbohydrate-active
catalytic proteins. Some glycosidases and glycanases have the
ability to catalyse transglycosylation in addition to hydrolysis;
however, extracting the data on such enzymes that ‘cut and
paste’ plant cell-wall polysaccharides can be laborious, especially when an initial search for what the user considers the
‘appropriate’ keyword fails. For example, ‘transglycosidase’,
‘transglycosylase’, or ‘glycosyltransferase’ have not been incorporated into the headline entries under ‘glycosidases’ but are
mentioned only marginally, if at all. On the other hand, many
databases provide entries which are not directly related to the
term sought, so users have to go through numerous categories
manually. Nevertheless, more than 19 databases that to some
extent provide information on plant wall-modifying enzymes
and/or their genes are currently available online (Table 1).
One of the most browsed enzyme databases is that of the
International Union of Biochemistry and Molecular Biology
(IUBMB), covering enzyme nomenclature, published online in
1992. This is a list of recommended names for enzyme activities, classified into classes and sub-classes according to Enzyme
Commission (EC) numbers. It is based on the enzymes’ specific
substrate preferences and is therefore guided by the reactions
they catalyse. As this concept does not take into account the
structural topology and stereochemical mechanistic features
of enzymes, it can be applied only to those proteins whose
functions have been biochemically identified.
The same principle (i.e. classification based on EC numbers) was employed in constructing the BRENDA and
IntEnz/ENZYME databases (Table 1), which, in addition to
nomenclature, provide detailed information about enzyme
features (such as kinetics, stability, substrate specificity, products formed, cofactors, subunits, etc.) and gene sequences,
respectively. Unlike many other databases, BRENDA is
highly searchable and enables immediate access to helpful
data on enzyme sources and localization, reaction and specificity, stability, and structure. However, despite BRENDA’s
virtues, there are a few cases in which the relevant records are
supported by inappropriate references or are misleading. For
example, the endo-1,5-α-l-arabinanase entry lists Arabidopsis
thaliana and Gossypium hirsutum under the ‘Organism’ item,
wrongly implying that α-arabinanase was detected in those
two species. In fact, α-arabinanase activity was never detected
either in Arabidopsis or Gossypium. Instead, a commercial
(non-plant) α-arabinanase was used by the cited authors to
digest Arabidopsis cytosolic heteroglycans (Fettke et al., 2006)
and cotton wall polysaccharides (Zheng and Mort, 2008).
To avoid any misinterpretation, users should cross-reference
to verify the accuracy of data extracted from any online
resources.
The Carbohydrate-Active enZyme database
As an alternative to the EC system, Carbohydrate-Active
enZyme (CAZy) classifies carbohydrate-acting enzymes and
carbohydrate-binding proteins into families based on their
sequence and structural folding features (Henrissat, 1991;
Henrissat and Bairoch, 1993). In contrast to the EC-IUBMB
classification, this structure-based system does not assign EC
numbers (i.e. the substrate specificity and the type of reaction being catalysed). Thus a single CAZy family can include
enzymes which act on various substrates. It also allows inclusion of proteins of unknown function, avoiding any premature
prediction of possible enzyme activity (Cantarel et al., 2009).
The CAZy classification is based on structurally related
catalytic and carbohydrate-binding domains of proteins and
comprises five classes: (i) glycosyl hydrolases (GHs, including
the enzymes discussed above as glycosidases and glycanases);
(ii) glycosyltransferases (GTs, principally those mentioned as
catalysing ‘irreversible’ transglycosylation); (iii) polysaccharide lyases (PLs); (iv) carbohydrate esterases (CEs); and (v)
carbohydrate-binding modules (CBMs).
Sometimes the folding of proteins provides a better basis
for classification than a simple one-dimensional sequence
and allows hierarchical categorization (clustering) of different families whose members seem to be structurally related.
This phenomenon can be observed in the class of glycanohydrolases, where some families are grouped into ‘clans’ (clan
GH-A to GH-N) defined by similarities in 3D structure (fold)
and a highly conserved catalytic domain and catalytic mechanism, despite differences in complete amino acid sequences
(Henrissat and Bairoch, 1996).
The CAZy database provides numerous options for searching through the individual categories such as family, organism, or protein name, EC number, and mechanism. Despite
the availability of these options, an initial search for required
data may fail after the relevant category has been selected. For
example, searching for enzyme activity defined by the protein
name ‘β-xylosidase’ will reveal 501 hits, but displaying only
10 results per page. Clicking on the next 10 records yields the
error message ‘résultats de la recherche’. A user would then
intuitively use another option, such as entering the EC number (in this case EC 3.2.1.37), which, however, provides the
same hits and the same problem. As an alternative, a user
can try the Glycoside Hydrolase (GH) family classification,
Plant cell-wall enzymes | 3527
Table 1. Synopsis of databases providing information on plant cell wall polysaccharide-modifying enzymes and/or their genes
Database name Description
Web link
Data
available on:
Notes
PS PE GO EF
IUBMB Enzyme
Nomenclature
Enzyme Commission (EC)
classification of enzymes by
the reactions they catalyse
Comprehensive enzyme
information system
www.chem.qmul.
–
ac.uk/iubmb/enzyme
+
–
–
Includes brief description of the enzyme-catalysed reaction
www.brendaenzymes.org
–
+
–
+
CAZy
Carbohydrate Active
enZyme database
www.cazy.org
+
+
–
–
IntEnz/ENZYME
Integrated relational enzyme
database/swissprot enzyme
nomenclature database
Sib bioinformatics resource
portal
Protein Data Bank database
www.ebi.ac.uk/
+
intenz; http://enzyme.
expasy.org
http://expasy.org
+
+
–
–
Summarizes detailed characteristics of enzymes/functional gene
products abstracted from the literature; enzymes classified according
to the EC list.
New sequence-based classification system introduced; enzymes
categorized into the families; information on sequences through the
GenBank and UniProtKB link
Describes type of enzyme based on the recommendations of the EC;
gene sequences and ontology provided through the link to UniProtKB
+
+
www.rcsb.org
+
+
+
Kyoto Encyclopedia of Genes
and Genomes database
European Bioinformatics
Institute databases
Annotated protein sequence
database
Hierarchical catalogue of
eukaryotic orthologues
National Center for
Biotechnology Information
database
Plant Proteome Database
for Arabidopsis thaliana and
maize (Zea mays)
www.kegg.jp
+
+
+
www.ebi.ac.uk/
services
www.uniprot.org
+
+
+
+
+
+
http://cegg.unige.ch/ +
orthodb4
www.ncbi.nlm.nih.
+
gov
-
+
+
+
http://ppdb.tc.cornell. +
edu
+
+
http://plantcyc.org
–
+
–
http://labs.plantbio.
cornell.edu/XTH
http://bioweb.ucr.
edu/Cellwall
+
–
–
CWN
Plant Metabolic Network
database
Database of XTHs from
Arabidopsis, tomato and rice
Cell-Wall Navigator database
+
–
–
AmiGO
Gene ontology database
http://amigo.geneon- +
tology.org
The Arabidopsis Information www.arabidopsis.org +
Resource database
–
+
–
+
PlantTribes
Floral Genome Project
database
http://fgp.bio.psu.
edu/tribedb/10_
genomes/index.pl
+
–
+
pDAWG
An integrated database for
plant cell-wall genes
–
+
GHATAbase
Glycosyl Hydrolase And
Transglycosylase Activity
database
http://csbl1.bmb.uga. +
edu/pDAWG/species.
php
http://www.
–
homepages.ed.ac.uk/
sfry/GHATAbase.html
+
–
+
Lists many scientific databases such as PROSIT, ENZYME, OMA
basic etc., easily searchable; provides access to software tools
–
Provides information on 3D structures and similarities, ligands,
methods, etc.
–
Consists of sub-databases categorized according to the information
available on systems, genomics and chemistry
–
Provides access to biological databases such as ENA, IntAct,
InterPro
–
Easily searchable, most information linked to the European
Nucleotide Archive (ENA)
–
Lists eukaryotic orthologous protein-coding genes; no record on
plant proteins but many on fungal carbohydrate-active enzymes
–
Supports access to a variety of enzyme and nucleotide databases,
genome-specific resources etc.; provides tools for sequence analysis
and 3D structure display.
+
Stores experimental data from proteome and mass spectrometry
basic analysis, curated information about protein function, protein properties and subcellular localization; predicted protein can be searched
for experimental information.
–
Contains curated information from the literature and computational analy­
ses about the genes, enzymes, compounds, reactions and pathways.
–
Web page proposes and standardizes the XTH nomenclature; a list of
new gene names with links to the NCBI sequences also provided.
–
Contains gene families that are involved in sugar substrate generation
and primary cell-wall metabolism; linked to sources of the complete
genome sequences of Arabidopsis thaliana and Oryza sativa and to
those of UniProt and NCBI.
–
Includes all manual gene product annotations and electronic annotations
from all databases other than UniProtKB; possibility to set up a filter.
–
Includes the complete genome sequence of Arabidopsis in addition
to gene product information, metabolism, genome maps, genetic and
physical markers and seed stocks.
–
A classification system for plant proteins based on cluster analyses
of the inferred proteomes of 9 sequenced angiosperms; includes
information about domains, traditional gene family names and unified
common terms
–
Contains 19 complete plant genomes including 12 from algae; linked
to the Pfam database (includes annotations and additional family
information); provides data on subcellular localization and phylogeny.
+
A list of individual enzyme activities for which evidence was obtained
in plant protein extracts; readily searchable; valuable resource for
selecting plant organs from whichto extract and study enzymes of
interest. Provides notes and comments about the reaction products.
BRENDA
ExPASy
PDB
KEGG
EMBL-EBI
UniProtKB
OrthoDB
NCBI
PPDB
PMN
XTH World
TAIR
EF, Enzyme features; GO, Gene ontology/annotations; PE, plant enzymes; PS, protein sequences.
3528 | Franková and Fry
under which the EC numbers are associated with individual
GH families. In the case of EC 3.2.1.37, ten GH families are
attributed to this enzyme activity and users are left with their
own manual search to find the necessary information, such
as the organism source. Each GH family contains entries categorized into archea, bacteria, eukaryota. The problem may
occur when users wish to find plant-specific entries that are
included simply under ‘eukaryota’. Based on structural similarities, each GH family can include not only proteins with
defined enzyme activity, but also peptide fragments and predicted or unknown proteins. These data are supported by references (available through NCBI or UniProtKB), which are
sometimes ‘unpublished data’ or ‘direct submissions’ (to the
online database). Even enzyme activities with defined name
(and/or EC number) are sometimes only ‘predicted’, so there
is no evidence at transcript or protein level for the corresponding protein. Such information can be misleading since a user
would intuitively trust an entry bearing a given enzyme name
or EC number, believing that enzyme’s existence has been
proven at the protein level (in vivo or in vitro). Nevertheless,
CAZy remains the only complex database on enzymes which
form, cleave, or reconstitute bonds in carbohydrates.
From the whole range of CAZy groups, approximately 22
families seem to be associated with enzymes that may postsynthetically modify the plant cell wall (Table 2; note that
families or enzyme activities that do not include any plant
member are omitted). Plant glycosidases (Fig. 2A, B) are
mostly grouped in GH families 1, 2, 3, 27, 29, 31, 35, 36,
38, 51, and 95, while plant glycanases (Fig. 2C) fall into GH
families 2, 5, 9, 10, 16, 17, 28, and 81. However, the boundaries between glycosidase and glycanases GH families is not
always strictly determined; for example, family GH2 includes
both exo- and endo-acting members. Dedicated reversible
transglycanases are restricted to family GH16. In contrast,
dedicated transglycosidases do not seem to exist in any GH
family, although families GH1 and 31 contain bifunctional
enzymes with both glycosidase and transglycosidase activity
(Table 2).
Transglycosylases or transglycosylating glycosyl
hydrolases?
Most enzyme databases use the term ‘glycosyltransferase’
for enzymes that catalyse the transfer of sugar residues, usually one at a time, from an activated donor substrate to a
specific acceptor substrate, forming a new glycosidic bond.
Such enzymes can also be described as aglycone-glycoside
synthases, oligosaccharide synthases, and polysaccharide
synthases. The donor substrate is usually a nucleoside diphospho- or monophospho-sugar or a sugar 1-phosphate (Lairson
et al., 2008; Palcic, 2011), and since the bond broken in the
donor is more energetic than the newly formed one, the transglycosylation reaction is usually essentially irreversible. The
reaction may affect the acceptor’s mass, solubility, transport,
and bioactivity (Ross et al., 2001).
A second group of enzymes catalysing (reversible) transglycosylation, which is not prominently distinguished in
enzyme databases and calls for an individual search (and a
lot of patience), comprises glycosyl hydrolases that also possess appreciable transglycosylation activity. Such transglycosidase and transglycanase activities are known mostly from
fungi and bacteria. If one searches for plant transglycanases
and transglycosidases (not NDP-sugar-dependent), one
would find fructan:fructan 1-fructosyltransferase (a transβ-fructanase) and sucrose:fructan 6-fructosyltransferase (a
trans-β-fructosidase), disproportionating enzyme (D-enzyme
or 4-α-glucanotransferase) and amylo-(1,4–1,6)-transglucosylase (branching or Q-enzyme, a trans-α-glucanase which converts amylose to amylopectin) (ExPASy, BRENDA). A more
intensive search for cell-wall-modifying transglycanases
and transglycosidases reveals entries on XET (xyloglucan
endotransglucosylase) and trans-β-mannanase (mannan
endotransglycosylase), mostly on the CAZy and BRENDA
servers. Both trans-β-mannanase and XET are classified
among glycosyl hydrolases (families GH5 and GH16, respectively; www.cazy.org) because their mechanism of action and
structural affiliations are different from those classed as glycosyltransferases (GT families). Other than XET and transβ-mannanase, no records on plant wall-related enzymes that
catalyse transglycosylation are yet available in online database
directories, so data on transglycosylation activities (discussed
further in the sections ‘Inverting matters’ and ‘Predicted
activities vs. biological roles of GH families’) are currently
available only in original articles.
Nevertheless, further new homo-transglycosidase and
homo- and hetero-transglycanase activities are being discovered in plants (Hrmova et al., 1998, 2006, 2007; Fry et al.,
2008a; Kosík et al., 2010; Franková and Fry, 2011) although
the sequences of the corresponding proteins are not always
known.
Mechanisms of enzymic hydrolysis and
transglycosylation
The enzyme-catalysed hydrolysis of glycosidic bonds can
take place via either of two reaction mechanisms: single- or
double-displacement. The single-displacement mechanism
proceeds in one step through an oxocarbenium ion-like transition state with the assistance of two carboxylic acids at the
active site (usually glutamic and/or aspartic acid; McCarter
and Withers, 1994). One carboxylic acid (acting as a catalytic
base) is required for nucleophilic attack on water, while the
second (acting as a catalytic acid) brings about the cleavage of the glycosidic bond (Koshland, 1953; Sinnott, 1990;
Withers, 2001). The result of such a mechanism is the inversion of anomeric configuration (e.g. bond cleaved = α-lfucosyl–R; initial products = β-l-fucose + R–OH), and this
defines ‘inverting glycosidases’. Regardless of the initial product formed, the sugar released (in aqueous solution) soon
ends up as an equilibrium mixture of, for example, α-l-fucose
and β-l-fucose, as a result of mutarotation.
The double-displacement mechanism is achieved in two
steps: (i) the formation of an intermediate containing a
glycosyl–­enzyme ester bond; and (ii) its hydrolysis (Sinnott,
1990; Davies and Henrissat, 1995). Both steps proceed via
an oxocarbenium ion-like transition state and also require
Plant cell-wall enzymes | 3529
Table 2. Distribution of plant cell-wall-remodelling enzymes in CAZy families
Data on plant enzymes were laboriously extracted from the very long list of plant, fungal, and animal CAZymes, all of which were included in
the one group ‘Eukaryota’. Unnamed/predicted protein products with unknown function and fragments are not included; families or enzyme
activities that do not include any plant member are omitted.
Family Catalytic Enzyme
domain activity
Mechanism Transglyco- Reference on
EC
sylation
transglycosylation number
activity
Plant sources of the protein with known
function/activity
GH1
β-Glucosidase
Retaining
+
β-Mannosidase
Retaining
ND
β-Mannosidase
Mannosylglycoprotein
endo-β-mannosidase
Retaining
ND
–
3.2.1.25
Arabidopsis thaliana, Avena sativa, Brassica
napus, Carapichea ipecuanha, Carica papaya,
Cicer arietinum, Consolida orientalis, Corbicula
japonica, Hordeum vulgare, Lotus japonicus,
Malus × domestica, Manihot esculenta,
Medicago truncatula, Olea europea, Oryza
sativa, Pinus contorta, Solanum lycopersicum,
Trifolium repens, Vitis vinifera, Zea mays
Hordeum vulgare, Oncidium Gower Ramsey,
Oryza sativa, Solanum lycopersicum
Brassica oleracea
Retaining
ND
–
3.2.1.152
Arabidopsis thaliana, Lilium longiflorum
β-Galactosidase
Retaining
ND
–
3.2.1.23
Arabidopsis thaliana
α-Arabinofuranosidase
/β-1,4-xylosidase
Retaining
ND
–
3.2.1.55
/3.2.1.37
β-1,4-Xylosidase
Retaining
ND
–
3.2.1.37
β-Glucosidase
Retaining
ND
–
3.2.1.21
Retaining
β-Glucosidase
(preferred substrates
are polysaccharides,
thus ‘exo-β-glucanase’)
Retaining
β-Mannanase,
ND
–
3.2.1.–
Actinidia deliciosa, Arabidopsis thaliana,
Solanum lycopersicum, Fragaria × ananassa,
Hordeum vulgare, Malus × domestica,
Medicago sativa ssp. varia, Medicago
truncatula, Pyrus pyrifolia, Raphanus sativus
Arabidopsis thaliana, Camellia sinensis,
Hordeum vulgare, Medicago truncatula,
Populus tremula × alba, Solanum lycopersicum,
Zea mays
Gossypium hirsutum, Nicotiana tabacum,
Tropaeolum majus
Hordeum vulgare
+
Hrmova et al. (2006); 2.4.1.Schröder et al. (2006) /2.4.1.78
GH2
(β/α)8
(β/α)8
GH3
GH5
(β/α)8
trans-β-mannanase
Crombie et al. (1998); 3.2.1.21
Opassiri et al. (2003)
3.2.1.25
GH9
(α /α)6
β-1,4-Glucanase
(cellulase)
Inverting
ND
–
3.2.1.4
GH10
(β/α)8
β-1,4-Xylanase,
trans-β-xylanase
Retaining
+.
Johnston et al. (2013) 3.2.1.8
Hordeum vulgare, Coffea arabica, Daucus
carota, Glycine max, Lactuca sativa, Solanum
lycopersicum
Arabidopsis thaliana, Brassica napus,
Capsella rubella, Capsicum annuum, Citrus
sinensis, Colocasia esculenta, Cucumis melo,
Dimocarpus longan, Fragaria × ananassa,
Glycine max, Gossypium barbadense,
Gossypium herbaceum, Gossypium hirsutum,
Hordeum vulgare, Malus × domestica
Mangifera inica, Medicago truncatula, Nicotiana
tabacum, Oryza officinalis, Oryza sativa, Persea
americana, Phaseolus vulgaris, Picea glauca,
Picea sitchensis, Pinus radiata, Pinus taeda,
Pisum sativum, Populus alba, Populus alba ×
grandidentata, Populus tremuloides, Prunus
persica, Pyrus communis, Saccharum hybrid
cultivar R570, Triticum aestivum, Vitis vinifera,
Sambucus nigra, Solanum lycopersicum,
Sorghum bicolor
Arabidopsis thaliana, Carica papaya, Hordeum
vulgare, Nicotiana tabacum, Oryza sativa, Zea
mays
3530 | Franková and Fry
Table 2. (Continued)
Family Catalytic Enzyme
domain activity
Mechanism Transglyco- Reference on
EC
sylation
transglycosylation number
activity
Plant sources of the protein with known
function/activity
GH16
Xyloglucan
endotransglucosylase
Retaining
+
Xu et al. (1995);
Campbell and
Braam, 1999)
2.4.1.207
Xyloglucan
endohydrolase
Retaining
+
3.2.1.151
β-1,3-Glucanase
Retaining
ND
De Silva et al. (1993);
Fanutti et al. (1993);
Tabuchi et al. (2001);
Baumann et al.
(2007); Zhu et al.
(2012)
–
Actinidia deliciosa, Annona cherimola,
Arabidopsis thaliana, Asparagus officinalis,
Beta vulgaris, Betula pendula, Brassica
oleracea var. botrytis, Brassica rapa, Capsicum
annuum, Carica papaya, Cenchrus americanus,
Chrysanthemum × morifolium, Dahlia pinnata,
Daucus carota, Fagus sylvatica, Festuca
pratensis, Gerbera hybrid cultivar, Hordeum
vulgare, Litchi chinensis, Medicago truncatula,
Musa acuminate, Nicotiana tabacum, Oryza
sativa, Pisum sativum, Populus euphratica, Pyrus
communis, Pyrus pyrifolia, Rosa chinensi, Shorea
parvifolia, Solanum lycopersicum, Striga asiatica,
Triticum aestivum, Vitis labrusca × vinifera
Arabidopsis thaliana, Tropaeolum majus, Vigna
angularis
3.2.1.39
‘Lichenase’
(MLG-specific
β-1,4-glucanase)
Retaining
ND
–
3.2.1.73
GH17
β-Jelly roll
(β/α)8
Arabidopsis thaliana, Atropa belladonna, Avena
sativa, Beta vulgaris ssp. vulgaris, Brassica rapa,
Cicer arietinum, Cichorium intybus × endivia,
Citrus clementina × reticulata, Citrus jambhiri,
Citrus sinensis, Glycine max, Gossypium
hirsutum, Hevea brasiliensis, Hordeum vulgare,
Medicago sativa, Musa acuminata, Musa
paradisiaca, Nicotiana tabacum, Olea europaea,
Oryza sativa, Phaseolus vulgaris, Pisum
sativum, Salix gilgiana, Solanum lycopersicum,
Solanum tuberosum, Triticum aestivum, Vitis
vinifera, Zea mays
Avena sativa, Hordeum vulgare, Nicitiana
plumbaginifolia, Oryza sativa, Triticum aestivum
GH 27
(β/α)8
α-Galactosidase
Retaining
ND
–
3.2.1.22
GH28
β-helix
α-Galacturonidase
(‘exopolygalacturonase’)
Galacturonanase
(‘endopolygalacturonase’,
pectinase)
Inverting
ND
–
3.2.1.67
Inverting
ND
–
3.2.1.15
α-1,3-Fucosidase,
α-1,4-Fucosidase
Retaining
ND
–
3.2.1.51
α-Glucosidase,
α-Xylosidase
Retaining
+
Sampedro et al.
(2010)
3.2.1.20
/3.2.1.-
Arabidopsis thaliana
α-Xylosidase
Retaining
ND
–
3.2.1.-
Oryza sativa, Tropaeolum majus
GH29
GH31
(β/α)8
Coffea arabica, Coffea canephora, Cucumis
sativus, Glycine max, Helianthus annuus, Oryza
sativa, Pisum sativum
Arabidopsis thaliana, Brassica rapa ssp.
campestris, Oryza brachyantha, Oryza
coarctata, Oryza minuta, Zea mays
Arabidopsis thaliana, Brassica napus, Brassica
rapa ssp. campestris, Carica papaya, Cucumis
melo, Daucus carota, Eucalyptus globulus,
Fragaria chiloensis, Fragaria × ananassa, Glycine
max, Gossypium barbadense, Gossypium
hirsutum, Hypericum perforatum, Lilium
longiflorum, Medicago sativa, Musa acuminata,
Nicotiana tabacum, Oncidium Gower Ramsey,
Oryza brachyantha, Platanus × acerifolia, Prunus
armeniaca, Prunus domestica ssp. insititia,
Prunus persica, Pyrus communis, Salix gilgiana,
Solanum lycopersicum, Vitis vinifera
Arabidopsis thaliana
Plant cell-wall enzymes | 3531
Table 2. (Continued)
Family Catalytic Enzyme
domain activity
Mechanism Transglyco- Reference on
EC
sylation
transglycosylation number
activity
Plant sources of the protein with known
function/activity
GH35
(β/α)8
β-Galactosidase
Retaining
(inferred)
ND
–
3.2.1.23
GH36
(β/α)8
α-Galactosidase
Retaining
ND
–
3.2.1.22
GH38
(β/α)7
α-Mannosidase
Retaining
ND
–
3.2.1.24
GH51
(β/α)8
α-Arabinofuranosidase
Retaining
ND
–
3.2.1.55
ND
–
3.2.1.55
/3.2.1.37
GH81
ND
α-Arabinofuranosidase, Retaining
β-xylosidase
Inverting
β-1,3-Glucanase
Arabidopsis thaliana, Asparagus officinalis,
Brassica oleracea, Capsicum annuum, Carica
papaya, Cicer arietinum, Citrus sinensis,
Coffea arabica, Fragaria × ananassa, Glycine
max, Gossypium hirsutum, Hordeum vulgare,
Mangifera indica, Nicotiana tabacum, Oryza
sativa, Persea americana, Petunia × hybrida,
Prunus persica, Pyrus communis, Pyrus
pyrifolia, Solanum lycopersicum, Triticum
monococcum, Vigna radiata, Vitis vinifera,
Ziziphus jujuba
Arabidopsis thaliana, Cucumis melo, Cucumis
sativus, Oryza sativa, Pisum sativum, Zea mays
Arabidopsis thaliana, Capsicum annuum,
Medicago truncatula, Oryza sativa
Arabidopsis thaliana, Carica papaya, Fragaria ×
ananassa,Gunnera manicata, Hordeum vulgare,
Malus × domestica, Medicago truncatula,
Prunus persica, Pyrus communis, Pyrus
pyrifolia, Solanum lycopersicum
Arabidopsis thaliana
GH95
(α/α)6
α-1,2-Fucosidase
Inverting
CE6
(α/β/α)Sandwich
Xylan acetylesterase
CE8
β-Helix
CE13
PL1
ND
–
3.2.1.39
Arabidopsis thaliana, Glycine max
ND
–
3.2.1.63
Deacetylation –
–
3.1.1.72
Arabidopsis thaliana, Lilium longiflorum, Oryza
sativa
Arabidopsis thaliana, Hordeum vulgare
Pectin methylesterase
Demethyl­
esterification
–
–
3.1.1.11
(α/β/α)Sandwich
Pectin acetylesterase
Deacetylation –
–
3.1.1.-
Parallel
β-helix
Pectate lyase
β-Elimination
–
–
4.2.2.2
ND, not determined.
Allium cepa, Arabidopsis halleri ssp. halleri,
Arabidopsis thaliana, Brassica napus, Brassica
oleracea, Brassica rapa ssp. pekinensis,
Capsicum annuum, Citrus sinensis, Fragaria
× ananassa, Linum usitatissimum, Lycoris
aurea, Medicago truncatula, Nicotiana
benthamiana, Nicotiana plumbaginifolia,
Nicotiana tabacum, Olea europea, Oncidium
Gower Ramsey, Oryza rufipogon, Oryza sativa,
Petunia integrifolia subsp. inflata, Phaseolus
vulgaris, Physcomitrella patens, Picea abies,
Pisum sativum, Populus tremula × tremuloides,
Prunus persica, Pyrus communis, Salix gilgiana,
Sesbania rostrata, Silene latifolia ssp. alba,
Solanum lycopersicum, Solanum tuberosum,
Vitis riparia, Vitis vinifera
Lactuca sativa, Litchi chinensis, Medicago
truncatula, Oryza sativa, Populus trichocarpa,
Vitis vinifera, Vigna radiata var. radiata, Sorghum
bicolor
Arabidopsis thaliana, Carica papaya, Dianthus
caryophyllus, Fragaria chiloensis, Gossypium
barbadense, Gossypium herbaceum,
Gossypium hirsutum, Gossypium raimondii,
Hevea brasiliensis, Lilium longiflorum, Malus ×
domestica, Mangifera indica, Medicago sativa,
Musa acuminate, Nicotiana tabacum, Populus
tremula × Populus tremuloides, Prunus persica,
Rosa × borboniana, Salix gilgiana, Solanum
lycopersicum, Zinnia violacea
3532 | Franková and Fry
two acidic amino acid residues—one acting as a nucleophile
and the second as an acid/base catalyst. In the first (glycosylation) step, a nucleophilic residue attacks the anomeric centre (defined in Fig. 3) allowing displacement of the aglycone
and formation of the glycosyl–enzyme complex. At the same
time, the carboxylic group (functioning as an acid catalyst)
protonates the glycosidic oxygen (defined in Fig. 3), cleaving
the original glycosidic bond in the substrate. The glycosyl–
enzyme ester bond is then hydrolysed by water in the second
(deglycosylation) step. The same carboxylic group (now acting as a base) deprotonates the water molecule, forming a new
–OH group. Enzymes operating via this double-displacement
mechanism are called ‘retaining’ as overall they maintain
the initial conformation at the anomeric carbon (e.g. bond
cleaved β-d-glucosyl–R; products β-d-glucose + R–OH).
Retaining hydrolases of interest in connection with plant
cell walls include those in CAZy families GH1–3, 5, 10, 16,
17, 27, 29, 31, 35, 36, 38, and 51 (Table 2). Inverting GH families of interest include GH9, 28, 81, and 95 (www.cazy.org).
Inverting matters
Unlike inverting hydrolases, which catalyse only hydrolysis,
some retaining glycosylhydrolases can also participate appreciably in transglycosylation reactions (Koshland, 1953; Sinnott,
1990; Scigelova et al., 1999; Moracci et al., 2001; Tramice
et al., 2007). This is due to their ability to form a glycosyl–
enzyme complex which can then be attacked by an acceptor
substrate other than water, generating a new glycosidic bond
instead of releasing a reducing sugar as hydrolysis product.
This knowledge has been applied to the design and development of biotechnologically improved enzymes. New mutants
of retaining glycosylhydrolases (including those from plants;
Hrmova et al., 2002; Hommalai et al., 2007; Piens et al.,
2007) were created by selective intervention in their ‘hydrolytic domain’ such as the replacement of a catalytic nucleophile (e.g. Glu231 or Glu235 of the barley β-1,3-glucanase and
Cellulomonas β-1,4-xylanase respectively) by an inert, nonnucleophilic residue (e.g. Ser, Gly, Ala, or Cys; Withers, 2001;
Hrmova et al., 2002; Kim et al., 2006). These modified glycosylhydrolases possessed hitherto unknown synthetic abilities, generating novel glycoconjugates, and have been termed
glycosynthases (Mackenzie et al., 1998; Moracci et al., 2001).
The mutant glycosynthase itself cannot form a glycosyl–
enzyme intermediate complex because it lacks a catalytic
nucleophile. However, the formation of a glycosyl–enzyme
intermediate can be mimicked by the use of glycosyl fluorides as donors, which possess an anomeric configuration
opposite to that of the natural substrate and a fluorine atom
as a good leaving group (comparable to the UDP in UDPglucose; Withers, 2001; Kim et al., 2006; Kang et al., 2007).
Oligosaccharide fluorides were successfully employed in the
synthesis of long oligosaccharides by mutant versions of
plant hydrolases such as rice β-glucosidase, Populus XET, and
barley β-1,3-d-glucanases (Hrmova et al., 2002; Hommalai
et al., 2007; Piens et al., 2007). Owing to the low cost of glycoconjugate synthesis, the application of glycosylhydrolases
modified by genetic engineering has been put into practice
and often prevails over the traditional chemical synthesis and
glycosyltransferase approach (the use of natural enzymes
and expensive sugar-nucleotides; Withers, 2001; Piens et al.,
2007). However, screens for natural plant hydrolases capable
of significant transglycosidase or transglycanase activity at
low substrate concentrations are also of interest as they are
accessible from a wide diversity of plant taxa and are relatively cheap. Thus, glycosynthases together with ‘cheap’ and
widespread native retaining hydrolases represent a powerful synthetic tool for preparing new compounds with possible application in the carbohydrate and pharmaceutical
industries.
Transglycosylation catalysed by retaining hydrolases may
in many cases be observed only at high acceptor substrate
concentrations, capable of competing with water (the ‘acceptor substrate’ in hydrolysis). Such transglycosylation is often
described as ‘mechanistic’, and may be denigrated if unphysiologically high substrate concentrations are required, or even
overlooked because hydrolysis (which is irreversible) will, in
the end, inevitably exceed reversible transglycosylation during prolonged enzyme assays by tapping off constituents of
the interconverting glycoside pool:
Of the 156 CAZy families, only 22 contain plant enzymes
that appear likely to post-synthetically modify plant cellwall polysaccharides (Table 2), namely families GH1, 2,
3, 5, 9, 10, 16, 17, 27, 28, 29, 31, 35, 36, 38, 51, 81, and 95,
CE6, 8, and 13, and PL1. As mentioned above in the section ‘Transglycosylases or transglycosylating glycosyl hydrolases?’, the CAZy database reports the transglycosylating
ability of only two plant ‘GH’ families: GH16 and GH5, represented by XET and trans-β-mannanase activities, respectively. The sequences and 3D structures of these two types
of wall-acting enzyme place them in GH (not GT) families,
but their activities are associated solely or primarily with
transglycosylation (Table 2). The ability to catalyse transglycosylation is also recorded for members of some fungal
and bacterial GH families but for no other plant GH families. However, some additional retaining plant GH CAZymes
listed in Table 2 have been found experimentally to catalyse
not only hydrolysis but also transglycosylation reactions in
the presence of moderate acceptor substrate concentrations,
e.g. 1–5 mM (Crombie et al., 1998; Opassiri et al., 2003;
Schröder et al., 2006; Sampedro et al., 2010; Johnston et al.,
2013), although this is not recorded in the CAZy database.
Such concentrations are on the verge of being low enough
to be considered ‘physiological’, and the enzymes involved
Plant cell-wall enzymes | 3533
might also perform transglycosylation within the wall matrix.
Examples include soyabean β-glucosidase, fenugreek endoβ-mannanases, and clover α-galactosidase, which catalyse
in-vitro transglycosylation reactions at ~0.2–40 mM acceptor
substrate concentrations (Williams et al., 1977; Coulombel
et al., 1981; Nari et al., 1983a,b). Moreover, recently reported
trans-β-xylosidase, trans-β-xylanase, trans-β-galactosidase,
and trans-α-xylosidase activities were detected with 0.5,
0.5, 1, and 0.016 mM oligosaccharide substrates respectively
(Franková and Fry, 2011, 2012a). These concentrations can
be regarded as low and close to (or even lower than) those
occurring in muro, since cellulose and hemicelluloses constitute about 20–30% and up to 20%, respectively, of the
primary cell-wall dry weight (Varner and Lin, 1989). The discovery of novel transglycosylase activities in plants suggests
(Popper and Fry, 2008) that hemicelluloses and pectins in the
wall matrix may not be linked only by non-covalent bonds,
as was assumed in earlier cell-wall models (Northcote, 1972;
Monro et al., 1976).
The plant cell wall itself is not equipped with either a pool
of activated substrates (e.g. NDP-sugars) or the enzymic
machinery (glycosyltransferases) to synthesize polysaccharides de novo (Schröder et al., 2009). Therefore, the manufacture of new glycosidic bonds within the cell wall (e.g. during
wall integration of new polymers, restructuring of existing
material, and bonding of polysaccharides to each other) can
only be accomplished by means of transglycosylation reactions. It might be only a matter of time before other transglycosylation activities are discovered in plants and accepted as
being non-‘mechanistic’ in the context of their possible biological roles in vivo.
Plant-centred, but no plant-specific, CAZy families
No ‘pioneer’ CAZy family found only in plants
All the GH and CE families that include plant cell-wallremodelling CAZymes also have representatives in archaea,
bacteria, viruses, protists, and animals. In other words,
there is no protein family containing only plant CAZymes.
However, regarding the metabolism of xyloglucan (which is
unique to plants), two enzyme activities (XET and XEH)
seem to be highly conserved in plants. Both activities fall into
family GH16, which also includes keratin-sulphate endo1,4-β-galactosidase (EC 3.2.1.103), endo-1,3-β-glucanase
(EC 3.2.1.39), endo-(1,3-1,4)-β-glucanase (‘cellulase’; EC
3.2.1.4), lichenase (EC 3.2.1.73), β-agarase (EC 3.2.1.81), and
κ-carrageenase (EC 3.2.1.83). Despite having the activities
listed above, the XTH branch of family GH16 can be regarded
as a plant-specific CAZy family, since XET- and XEH-active
GH16 members are found only in plants. Nevertheless, a
few members of other families (mainly GH5, 7, 12, 44 and
74) show XEH activity (Gilbert et al., 2008; Vlasenko et al.,
2010; Ariza et al., 2011). These 1,4-β-glucan-degrading
enzymes (called ‘cellulases’, albeit sometimes highly xyloglucan-specific) are bacterial or fungal and their possible ability
to catalyse transfer with xyloglucan was not reported. Other
CAZy families which are best represented among plant wallrelated enzymes are the carbohydrate esterase families CE8
and CE13. The sequences falling into these two families were
predicted pectin methylesterases and pectin acetylesterases.
The importance of pectin methylesterase (PME) in planta is
implied by the fact that Arabidopsis contains at least 79 putative PME genes (Markovič and Janeček, 2004). For example,
the CAZy database lists about 65 records for Arabidopsis
PMEs and 2 for fragments thereof (www.cazy.org). Even
though all these sequences are included in the PME family,
only a few of them have been screened for a functional PME
product and possible biophysicochemical properties (for
details see Richard et al., 1994, 1996; Francis et al., 2006).
Likewise, out of 66 entries for Arabidopsis available on the
UniProtKB server, 14 represent ‘putative’ and 44 ‘probable’
PMEs, indicating that the actual enzyme activity was tested
in very few cases.
Predicted activities vs. biological roles of GH families
Both the spatial and temporal regulation of gene expression
and variation in enzyme activities are routinely monitored
by: (i) quantification of mRNAs (by in-situ hybridization
and gene-specific microarrays); (ii) measurement of protein
steady-state levels; (iii) tissue printing (hybridization, immunohistochemistry); and (iv) the study of mutants with overexpressed or knocked-down genes. Sometimes mRNA levels
do not faithfully predict actual enzyme activities (because
there may be post-transcriptional regulation), which makes
the interpretation of biological roles more difficult. Likewise,
overexpressing or silencing of selected genes may not produce
any morphological phenotype in transgenic plants, especially
when the gained/lost function can be compensated for by
non-affected isoforms or other regulatory mechanisms. In
such cases, a reliable ‘wet biochemical’ approach (e.g. sensitive enzyme assays, histochemistry, polysaccharide content
and composition analysis, reducing sugar assays) becomes an
indispensable complement to functional genomics.
The activity of an archetypal plant transglycanase – XET
– is associated with various key biological functions (cellwall loosening and expansion, fruit softening, germination,
reserve mobilization, secondary wall deposition, wall assembly and strengthening) and is attributed to multiple isoforms.
For example, the Glycine max, Arabidopsis lyrata, A. thaliana,
and Zea mays genomes are equipped with 64, 39, 33, and 32
XTH genes, respectively, that were predicted to encode functional gene products (Michel et al., 2001; Nishitani, 2005;
Eklöf and Brumer, 2010). Based on the available genome
data, the XTH enzymes and their genes have been grouped
into a phylogenetic tree which is divided into three clades (I,
II, and IIIb) expected to exhibit only XET activity (Eklöf and
Brumer, 2010), and a fourth (IIIa) assumed to work predominantly as XEH. Their ability to fulfil this role will discussed
in the section ‘Xyloglucan endotransglucosylase/hydrolase’.
Apart from the action of ‘dedicated’ transglycosylases
(XTHs with XET activity), many other ‘part-time’ transglycosylation activities have been reported in plants. So far, only
six plant glycosyl hydrolases with known protein sequences
have been demonstrated to catalyse glycosyl transfer: rice
Bglu β-glucosidase (Opassiri et al., 2003) and nasturtium
3534 | Franková and Fry
β-glucosidase (Crombie et al., 1998), both showing trans-βglucosidase activity; Arabidopsis AtXyl1 α-xylosidase (with
trans-α-xylosidase activity; Sampedro et al., 2010), barley
HvMAN1 β-mannanase and tomato LeMan4 mannanase
(with trans-β-mannanase activity; Hrmova et al., 2006;
Schröder et al., 2006); and papaya CpaEXY1 β-xylanase
(with trans-β-xylanase activity; Johnston et al., 2013). Their
function, regulation, and activity were examined not only at
the transcriptional level but also at the protein level (enzyme
assays and reaction product analysis). Based on structural
features, they were all predicted to function as glycosyl
hydrolases (Table 2). However, the importance of ‘cutting
and pasting’ glycosidic bonds in vitro and its relevance in vivo
was highlighted. Fig. 4 depicts proposed biological functions
of plant transglycosylases, including those whose protein
sequence is not known (section ‘Inverting matters’).
Gene expression does not guarantee
enzyme action in vivo
Newly discovered enzyme activities call for subsequent protein purification and genetic studies which might reveal the
identity of proteins responsible for the reactions catalysed.
Equally, the discovery of expressed genes calls for enzymological and biological studies of the reactions which their
translation products may catalyse. The finding that a given
gene is transcribed such that its mRNA can be detected, via
cDNA analysis, in a particular cell at a particular time, and
that the gene in question fits in a particular CAZy class, does
not prove that that gene’s product is capable of catalysing the
CAZy-predicted reaction, still less that it actually does so in
vivo. These issues require testing experimentally.
There is an important distinction between enzyme activity
(e.g. assayed in vitro, under optimized conditions, with substrates arbitrarily chosen by the experimenter) and enzyme
action (as occurring in muro, with natural substrates).
A protein might fail to exhibit its CAZy-predicted activity in vitro for any of several reasons, such as: (i) badly chosen conditions (pH, ionic strength, cofactor availability), not
accurately mimicking those occurring in vivo; (ii) enzyme
denaturation during extraction; (iii) heterologously produced protein (e.g. in Escherichia coli or Pichia) lacking correct post-translational modifications, e.g. N-glycosylation;
and (iv) the true substrate of the enzyme may not be as predicted by CAZy. This last uncertainty can readily be explored
by assays on pure (e.g. heterologously expressed) protein.
In many cases, the enzyme is indeed active, although there
are exceptions. For example the protein encoded by a putative plant α-fucosidase gene showed no α-fucosidase activity in vitro (Tarragó et al., 2003). As a second example,
plant bifunctional α-arabinofuranosidase/β-xylosidase (e.g.
MsXyl1 of Medicago, ARA-I/XYL of barley, and AtBXL1
of Arabidopsis; Lee et al., 2003; Xiong et al., 2007; Arsovski
et al., 2009; see section ‘The mutation/RNAi approach’)
is placed in CAZy family GH3, which mainly contains
β-glucosidases, β-glucanases and β-xylosidases. In this case,
the unexpected α-arabinofuranosidase activity could have
been easily overlooked as plant α-arabinosidases belong to
GH51. A third example, Arabidopsis AtFuc1 α-fucosidase,
was primarily thought to be acting on α-1,2 fucosyl linkages
other than those of xyloglucan (de la Torre et al., 2002). Later,
it was reported that AtFuc1 hydrolyses both 3- and 4-linked
fucoses but not 2-linked α-fucose nor the α-fucose that is
1,3-linked to the innermost GlcNAc residue of glycoproteins
(Zeleny et al., 2006). Thus the enzyme was moved from family
GH95 to GH29. The ‘true’ α-1,2-fucosidase (GH95) associated with xyloglucan metabolism is encoded by AtFXG1 (also
AXY8 or AtFuc95A) and exhibits activity against XXFG and
2′-fucosyllactose (de la Torre et al., 2002; Günl et al., 2011).
Like AtFuc1, it is inactive on pNP-α-Fuc.
More worryingly, even if a protein does possess a given
enzymic activity on soluble substrates in vitro, it might fail
to exert any corresponding action in vivo owing to multiple
reasons such as: (i) the endogenous substrate may be inaccessible to endogenous enzyme, for example because the putative
substrate (e.g. mannan) is shrouded by some other polysaccharide (e.g. pectin) (‘masking’; Marcus et al., 2010); (ii) the
enzyme may be localized in different cells, or in different parts
of the cell, from the putative polysaccharide substrate; (iii)
the apoplast of the cells in question may have an inappropriate pH for action of the enzyme; (iv) activators, for example
cofactors, may be absent in vivo; and (v) inhibitors may be
present in vivo.
For any of these reasons, in-vitro assays of enzyme action
may not correctly predict in-vivo action, and one may be led
to false conclusions about the biological role of a gene. A successful combination of classical ‘wet’ biochemistry, applied
both in vitro and in vivo, plus a functional genomic approach
may bring new and reliable insights into the roles of polysaccharide-modifying enzymes in plants, as will be discussed further in the section ‘Experimentally investigating wall enzyme
action in vivo’.
Assaying polysaccharide-restructuring
enzyme activities
Numerous enzymes are extractable from plant cell walls in
aqueous buffers (sometimes assisted by high salt), including glycanases, glycosidases, esterases, proteinases, transglycanases, transglycosidases, transacylases, peroxidases,
oxidases, and lyases. Often, their substrate specificities suggest physiological significance in modifying wall components
(Minic, 2008). Here are discussed methods for assaying glycanases, glycosidases, transglycanases, and transglycosidases.
Glycanases and glycosidases
Glycanase and glycosidase activities are assayed on substrates
in which a glycosidic bond is hydrolysed. Methods for glycanase-catalysed endo-hydrolysis of polysaccharides include
(in approximate order of decreasing sensitivity):
(i) Loss of a polysaccharide solution’s viscosity, measured
in a simple viscometer (Farkaš and Maclachlan, 1988).
Plant cell-wall enzymes | 3535
Polysaccharides are also cleaved non-enzymically by ascorbate-generated hydroxyl radicals (•OH) (Fry, 1998); therefore,
in these highly sensitive assays, enzyme extracts should be
freed of ascorbate etc., for example by dialysis.
(ii) Increase in the number of reducing termini (oxo groups,
assayed colorimetrically). It should be checked that the new
‘reducing termini’ formed are not monosaccharides released
by glycosidases, nor mid-chain oxo groups introduced by •OH
(Fry et al., 2001; Vreeburg and Fry, 2005).
(iii) Release of radioactive oligosaccharides from a reducingend-labelled polysaccharide, e.g. [galactitol-3H]galactan (Fry,
1983) or [glucitol-3H]xyloglucan (Zhu et al., 2012). Since most
glycosidases attack at the non-reducing end, this method is
unlikely to be compromised by contaminating glycosidases.
(iv) Increase in products insoluble in a precipitant (e.g. ethanol). For convenience, the substrate can be prelabelled with
fluorescent or coloured tags (as in azo-xylan).
(v) Release of water-soluble dyed products from artificially cross-linked polysaccharides (e.g. azurine-crosslinked
polysaccharides).
Glycanases and glycosidases can also be assayed on fully
defined oligosaccharides (Fig. 2A–C) that model biologically
relevant polysaccharides. Products generated are analysed by
TLC or HPLC: monosaccharides are diagnostic of glycosidase
activities; oligosaccharides smaller than the starting material
but unaccompanied by monosaccharides indicate glycanases.
Alternative assays for certain glycosidases employ fluorogenic or chromogenic model substrates (e.g. p-nitrophenyl or
4-methylumbelliferyl β-galactoside; NP- and 4-MU-β-Gal,
respectively). However, such substrates are not always suitable: plant extracts that efficiently hydrolyse XGOs do not recognize NP-α-Xyl or NP-α-Fuc (Fanutti et al., 1991; Léonard
et al., 2008).
Transglycanases and transglycosidases
In transglycosylation
products may be
reactions, the reactants and
indistinguishable, for example
(where , ,
and are chemically identical sugar residues as, for example,
depicted for homo-transglycanases in Fig. 4), so some kind of
labelling is often used. For transglycanases, the acceptor substrate can often (perhaps always) be an oligosaccharide, even
if the donor must be a polysaccharide. A radiochemically or
fluorescently tagged oligosaccharide is incubated with a nonlabelled polysaccharide and the diagnostic reaction product is
recognized by its label and large size. Sensitive labels include
tritium (3H), sulphorhodamine (SR), anthranilic acid (AA),
and fluorescein isothiocyanate (FITC). Some transglycanases
discriminate between differently labelled acceptor substrates:
for example, a Tropaeolum XTH has Km values for [3H]XLLG,
XLLG–SR, XLLG–FITC, and XLLG–AA of 60, 81, 130, and
530 μM respectively, indicating a low affinity for XLLG–AA;
in contrast, turnover numbers of the enzyme for the same substrates are respectively 20, 400, 1300, and 79 molecules of substrate per molecule of enzyme per hour (Kosík et al., 2011).
Thus absolute reaction rates measured on fluorescent substrates may be misleading, albeit useful for screening purposes.
The labelled high-Mr ‘hybrid’ product formed in such
transglycanase assays is separated from remaining unreacted
acceptor by a size-dependent method, for example paperbinding (if the acceptor, e.g. [3H]XXXGol, is easily washed
off paper whereas the donor, e.g. xyloglucan or MLG, has
an affinity for cellulose and remains bound), paper chromatography, gel-permeation chromatography, and ethanol
precipitation. With paper chromatography, all polysaccharides will remain immobile in solvent mixtures such as ethyl
acetate/acetic acid/water, while many oligosaccharides tested
as acceptors migrate satisfactorily away from the origin;
however, cello-, xylo-, manno- and MLG-oligosaccharides
(MLGOs) remain partially or completely at the origin owing
to their own affinity for cellulose, so paper chromatography is
not recommended for these.
Transglycosidases transfer only a single sugar residue, so a
labelled acceptor substrate would not increase greatly in size.
A convenient alternative is dual-labelling: a donor substrate radiolabelled in an appropriate sugar residue ( ) reacts with an acceptor substrate that is physically separable, for example by virtue of
a cationic ‘label’ (⊕). The product of interest carries both labels:
As with hydrolases, transglycosylases can be assayed on a nonlabelled oligosaccharide (as both donor and acceptor), e.g. by TLC
analysis. The products proving transglycosylation are those larger
than the substrate; smaller products are less informative because
they could be either hydrolysis products or leaving groups formed
by transglycosylation. Fluorescently and radiolabelled products
can be also assayed by capillary electrophoresis and high-voltage
paper electrophoresis respectively. Other techniques such as HPLC,
nuclear magnetic resonance spectroscopy, or matrix-assisted laser
desorption ionization–time-of-flight (MALDI–TOF) mass-spectrometry are an alternative to a quick semi-quantitative TLC, but
not applicable when radiolabelled substrates are used.
Specific examples of polysacchariderestructuring activities
Hydrolases
Plant cell walls contain more than 12 glycosidase activities (e.g. β-glucosidase, β-galactosidase, β-xylosidase,
α-xylosidase) and more than nine glycanases (e.g.
β-mannanase, β-xylanase) (reviewed by Labavitch, 1981;
3536 | Franková and Fry
Fischer and Bennet, 1991; Hrmova and Fincher, 2001; Fry,
2004; Libertini et al., 2004; Baumann et al., 2007; Gilbert
et al., 2008; Minic, 2008; Schröder et al., 2009). We recently
surveyed 57 species for such activities using simple oligosaccharides that model cell-wall polysaccharides (Franková
and Fry, 2011). The extensive results (exemplified in
Table 3) are available in GHATAbase (Table 1). Tables 4
and 5 give fuller lists of plant glycanases and glycosidases,
respectively, potentially attacking wall polysaccharides
(Labavitch, 1981; Fry, 1995, 2004; de la Torre et al., 2002;
Minic, 2008). Known activities are, together, theoretically
capable of hydrolysing most of the major glycosidic bonds
in wall polysaccharides (except RG-II). This, however, is
not to imply that all, or any, of them are present in vivo
at sufficiently high activity to completely lyse the wall.
Nevertheless, the numerous hydrolase activities certainly
contribute to the diverse examples of wall restructuring
occurring during normal plant growth and development, as
already discussed.
Transglycanases
Trans-β-xylanase
The GHATAbase study, using fully defined oligosaccharide
substrates, revealed not only hydrolases but also transglycanase and transglycosidase activities, several of which were
new (Franková and Fry, 2011). For example, trans-β-xylanase
activity converts Xyl6 to Xyl9 plus Xyl3:
(Johnston et al., 2013). Here the donor was high-Mr xylan,
the acceptor was [3H]Xyl5-ol, and the large radioactive
‘hybrid product’ was recognized by its immobility on paper
chromatography (albeit slightly contaminated by unreacted
[3H]Xyl5-ol). The authors attributed the trans-β-xylanase
activity to a protein previously characterized as β-xylanase;
thus the same protein can catalyse both endo-hydrolysis and
endo-transglycosylation, the ratio between these depending
on the acceptor-substrate concentration.
Xyloglucan endotransglucosylase/hydrolase (XTH)
Some transglycanases are undetectable on simple, well-defined
oligosaccharides because the donor needs to be a polysaccharide. The first in-vitro demonstrations of XET activity were
in plant enzyme extracts incubated with xyloglucan (donor)
plus a labelled XGO (acceptor), for example [3H]XXFG
(Fry et al., 1992) or XGO–PA (pyridylamino; Nishitani and
Tominaga, 1992). The reaction generated a large polysaccharide–XGO conjugate. The enzymes had high affinity for
XXFG (Km 50 μM) and higher affinity for XXXG (Km 33 μM)
and XLLG (Km 19 μM). Polysaccharides other than xyloglucan had little, if any, donor ability (Fry et al., 1992). Nishitani
and Tominaga (1992) showed that the enzyme required the
donor to be a polysaccharide of Mr >10 000 for appreciable
activity. Certain XTHs can, however, utilize oligosaccharides
as both donor and acceptor: for example:
XXX ′ XXX + XXXXXX ↔ XXXXXXXXX + XXX ( ′ indicates the cleaved bond );
whereas trans-β-xylosidase activity transfers one residue at a time:
X ′XXXXX + XXXXXX ↔ XXXXXXX + XXXXX
In a crude extract, these two activities are distinguishable
by their largest products: trans-β-xylanase immediately starts
making Xyl9, whereas trans-β-xylosidase initially produces
Xyl7 and would not begin to make Xyl9, if any, until after a lag
period during which Xyl7 and Xyl8 were sequentially generated.
An alternative assay, modelled on one widely used for
XET, also revealed trans-β-xylanase activity in plant extracts
where is a fluorescent label (Saura-Valls et al., 2006), albeit with
low affinity (Km 0.4 mM for donor and 1.9 mM for acceptor).
Besides assays on extracted plant proteins, a genomic
approach can be adopted. Plants have numerous ‘cell-wall
genes’ (Mao et al., 2009): Arabidopsis has 730 open reading
frames encoding putative glycosyltransferases and -hydrolases (Henrissat et al., 2001), 33 of which are XTHs. Work
is in progress to test experimentally the predicted activities
of encoded XTH-like proteins produced heterologously,
e.g. in the yeast Pichia, and the following Arabidopsis XTHs
do exhibit XET activity in vitro: XTH22 (formerly TCH4;
Purugganan et al., 1997), 14, and 26 (Maris et al., 2009); and
Table 3. Some hydrolase and transglycosylase activities detected in a survey of land plants, assayed on well-defined model substrates
Substrate cleaved
Bond broken
Glycosidase (exo)
Glycanase (endo)
Transglycosidase (exo)Transglycanase (endo)
Mannohexaose
β-Man
β-Mannosidase (+)
α-Ara
α-Arabinosidase (+)
β-Mannanase (±)
?
–
Arabinohexaose
Xylohexaose
β-Xyl
β-Xylosidase (+)
α-Xyl
α-Xylosidase (+)
β-Xylanase (+)
NA
Trans-β-xylosidase (+)
XXXG
α-Fuc
XLXG and 1st Gal of XLXG β-Gal
α-Fucosidase (+)
NA
β-Galactosidase (+)
NA
XXLG
β-Galactosidase (±)
NA
XXFG
β-Gal
–
Trans-α-arabinosidase (±) Trans-α-arabinanase (±)
Trans-α-xylosidase (+)
–
Trans-β-xylanase (+)
NA
NA
Trans-β-galactosidase (+) NA
Trans-β-galactosidase (+) NA
+, activity detected in most or all land plants; ±, activity detected in few land plants; –, activity not detected in land plants;?, data uncertain;
NA, not applicable. Data from Franková and Fry (2011).
Plant cell-wall enzymes | 3537
Table 4. Range of cell-wall-related glycanase activities reported in plant extracts
Cleaved bond (bold, underlined)
Polysaccharide whose backbone Name for the enzyme activity
could potentially be hydrolysed
...(1→4)-α-GalA-(1→4)-α-GalA-(1→4)... Pectic homogalacturonan
...(1→4)-β-Gal-(1→4)-β-Gal-(1→4)...
...(1→4)-β-Glc-(1→4)-β-Glc-(1→4)...
Pectinase, galacturonanase,
endo-polygalacturonase
Galactanase
Galactan/arabinogalactan domains of
RG-I
Cellulose, MLG; sometimes xyloglucan Cellulase, β-1,4-glucanase
Reference
Taylor et al. (1993); Ghiani et al. (2011)
Lazan et al. (2004)
Truelsen and Wyndaele (1991); Ohmiya
et al. (1995)
...(1→4)-β-Glc-(1→4)-β-Glc-(1→3)...
...(1→3)-β-Glc-(1→4)-β-Glc-(1→4)...
MLG
...(1→4)-β-Glc-(1→4)-β-Glc-(1→4)...
with α-Xyl on O-6 of 2nd Glc
Xyloglucan
...(1→3)-β-Glc-(1→3)-β-Glc-(1→3)...
(with β-Glc on O-6 in laminarin)
Callose, laminarin
...(1→4)-β-Man-(1→4)-β-Man-(1→4)... Mannan
‘Lichenase’ (MLG-specific
β-1,4-glucanase)
Xyloglucan endo-hydrolase (XEH),
xyloglucan endo-glucanase (XEG)
Hrmova and Fincher (1993); Martin and
Somers (2004)
Mannanase
Dahal et al. (1997); Schröder et al.
(2006)
Ronen et al. (1991); Johnston et al.
(2013)
Cota et al. (2007); Mizuno et al. (2008)
Xylan
Xylanase
...(1→4)-β-GlcNAc-(1→4)-βGlcNAc-(1→4)...
Chitin (of pathogenic organism)
Chitinase (defence-induced)
XTH12, 13, 17, 18, and 19 (Maris et al., 2011). In addition,
XET activity has been confirmed for: ZmXTH1 of maize
(Genovesi et al., 2008); PttXET16A of poplar (Johansson
et al., 2004); HvXET5 of barley (Hrmova et al., 2007); and
BRU1 of soyabean (Oh et al., 1998).
All putative XTH translation-products tested to date have
exhibited XET activity with tamarind xyloglucan as donor.
Relatively subtle differences in substrate preference exist,
e.g. between fucosylated and non-fucosylated xyloglucans
(Purugganan et al., 1997) and between contrasting acceptor XGOs (Maris et al., 2009). More pronounced variation
in pH preference was found: AtXTH12 and AtXTH17 have
pH optima of 5.0 and 7.5 respectively (Maris et al., 2011),
possibly indicating physiological differences in role. However,
the main differences between XTHs appear to be in the genes’
promoters, such that XET activity can be induced in different tissues and in response to different environmental stimuli
(Nishitani, 2005; Becnel et al., 2006).
Among the group IIIa XTHs, examples from
Tropaeolum (Fanutti et al., 1996; Baumann et al., 2007),
Vigna (Tabuchi et al., 2001) and Arabidopsis (AtXTH31;
Zhu et al., 2012; Kaewthai et al., 2013) are the only gene
products for which endo-hydrolytic activity has so far been
demonstrated in addition to low or very low XET activity.
Two Tropaeolum XTHs, TmNXG1 and TmNXG2, produced in the seed after germination, catalyse both hydrolysis (XEH activity) and transglycosylation (XET activity,
especially at higher substrate concentrations) of xyloglucan (Fanutti et al., 1996). Hydrolysis may fulfil the need
to mobilize seed-reserve xyloglucans for the nutrition of
the young seedling. Structural features indicate that these
two enzymes acquired XEH by loss-of-function mutations
Tabuchi et al. (2001); Zhu et al. (2012)
Laminarinase, β-1,3-glucanase
...(1→4)-β-Xyl-(1→4)-β-Xyl-(1→4)...
...(1→4)-β-GlcN-(1→4)-β-GlcN-(1→4)... Chitosan
Hrmova and Fincher (2001)
Chitosanase (constitutively present)
Ouakfaoui and Asselin (1992); Hung
et al. (2002)
of an XET-active ancestor (Baumann et al., 2007). By
comparison with these features, two of the 33 Arabidopsis
XTH genes (XTH31 and XTH32) were predicted to
encode XEH-active enzymes. Recently, this prediction
has been verified for XTH31 (AtXTH31; Zhu et al., 2012;
Kaewthai et al., 2013). When produced in Pichia cells,
XTH31 exhibited very slight XET activity, but >5000-fold
greater XEH activity (Zhu et al., 2012).
Trans-β-mannanase
In tomato fruit, trans-β-mannanase activity was detected with
high-Mr mannan as donor and [3H]manno-oligosaccharides
as acceptor (Schröder et al., 2006). The activity was attributed
to a previously characterized β-mannanase protein which
even at low acceptor substrate concentrations (0.18 mM) also
effected transglycosylation. Since this enzyme’s β-mannanase
activity can be assayed on Man6, it would be expected that
its trans-β-mannanase activity would also be detectable
with Man6 as donor (and acceptor) substrate. However, the
GHATAbase survey revealed no trans-β-mannanase activity
on 1.6 mM Man6 in any of the 57 extracts studied. Although
the list of 57 did not include tomato, this observation may
indicate that trans-β-mannanase activity is not widespread in
the plant kingdom even though β-mannosidase is (Franková
and Fry, 2011) and β-mannanase was found to be evolutionarily ancient and involved in diverse biological processes
(Yuan et al., 2007).
MLG:xyloglucan endotransglucosylase (MXE)
Several studies have explored the possibility that non-xyloglucan polysaccharides might serve as donors in conjunction
with XGOs as acceptors—‘hetero’-transglycanase activities.
Hrmova et al. (2007) tested the substrate specificity of a purified
3538 | Franková and Fry
Table 5. Range of cell-wall-related glycosidase activities reported in plant extracts
Non-reducing terminal
residue cleaved
Polysaccharide possessing Polysaccharide possessing Names for the
such termini (backbone)
such termini (side chains) enzyme activity
Reference
α-Araf-(1→5...
Pectic arabinan/arabinogalactan Pectic arabinan and
α-Arabinosidase
arabinogalactan, arabinoxylans,
and glucuronoarabinoxylans
Lee et al. (2003); Rosli et al.
(2009)
α-Fuc-(1→2...
–
α-Fuc-(1→3...
–
Léonard et al. (2008); Franková
and Fry (2011); Günl et al. (2011)
de la Torre et al. (2002); Zeleny
et al. (2006)
α-Araf-(1→3...
F of XXFG at non-reducing
terminus of xyloglucan
Wall glycoproteins and
glycolipids?
α-1,2-Fucosidase
α-1,3–1,4-Fucosidase
α-Fuc-(1→4...
α-Gal-(1→6...
–
Galactomannans/
galactoglucomannans
α-Galactosidase
Appukuttan and Basu (1987)
β-Gal-(1→4...
Galactan/arabinogalactan
domains of RG-I
Pectic β-galactan and
arabinogalactan domains of
RG-I, some xylans
β-Galactosidase
De-Veau et al. (1993); Kaneko
and Kobayashi (2003)
Xyloglucan
β-Galactosidase (widespread
activity)
de Alcântara et al. (1999);
Sampedro et al. (2012)
Xyloglucan
β-Galactosidase (rare activity)
Buckeridge et al. (1997);
Franková and Fry (2011)
Pressey and Reger (1989);
García-Romera and Fry (1994);
Tanaka et al. (2002)
Crombie et al. (1998); Hrmova
et al. (1996); Opassiri et al. (2003)
β-Gal-(1→3...
β-Gal-(1→2... in XLXG and 1st –
L of XLLG
β-Gal-(1→2... in XXLG
α-GalA-(1→4...
Pectic homogalacturonan of
DP ≥ 5
β-Glc-(1→4...
Cellulose, MLG, xyloglucan,
callose, laminarin
Laminarin
β-Glucosidase,
‘exo-β-glucanase’
Mannans
–
β-Mannosidase
Xylans
Single Xyl at non-reducing termi-α-Xylosidase
nus of xyloglucan
Xylans (xylogalacturonans?)
β-Xylosidase
α-Galacturonidase,
‘exo-polygalacturonase’
β-Glc-(1→3...
β-Glc-(1→6...
β-Man(1→4...
α-Xyl-(1→6...
β-Xyl-(1→4...
Hrmova et al. (2006); Franková
and Fry (2011)
Fanutti et al. (1991); Günl and
Pauly (2011)
Martínez et al. (2004); Minic et al.
(2004); Franková and Fry (2011)
β-Xyl-(1→3?..
barley XTH (HvXET5). With XGO–SR as acceptor, the preferred donor was xyloglucan, but certain other water-soluble,
substituted (1→4)-β-d-glucans were also effective donors: reaction rates relative to that with xyloglucan as donor (rate ‘100’)
were hydroxyethylcellulose, 44; sulphocellulose, 5; and carboxy­
methylcellulose, 0.4. Probably the hydroxyethyl ether and sulphate ester groups (both mainly linked to O-6 of β-Glc residues)
sufficiently resembled the 6-O-linked α-Xyl residues of xyloglucan to fit in the enzyme’s donor site. Water-soluble β-glucans
with different backbone linkages were much less effective: e.g.
barley MLG, relative rate 0.2; Cetraria MLG (‘lichenan’), 0;
(1→3)-β-glucan, 0; and glucomannan, 0. Furthermore, this
purified XTH was 1000-times more effective with XGO–SR as
acceptor substrate than with cello-oligosaccharide–SRs; it thus
has a very high but not absolute specificity for xyloglucan and
certain artificial (1→4)-β-glucans with hydrophilic side chains,
whereas its ‘MXE’ reaction rate (with MLG as donor substrate)
is 0–0.2% of its XET rate. Hrmova et al. (2007) suggested that
other, untested XTHs might exhibit higher MXE activity.
A good test of this suggestion is to assay crude plant
extracts, which will contain multiple isoenzymes. In fact,
Ait Mohand and Farkaš (2006) had conducted such assays
in Tropaeolum extracts. In agreement with the HvXET5
data, they showed that the XET donor preferences were
xyloglucan > HEC > CMC. They also detected heterotransglycanase activities (donor, xyloglucan; acceptor
cello- or laminari-oligosaccharide–SRs, these acceptors
lacking side chains) and suggested that the results indicate the plant’s ability to covalently link polysaccharides
to qualitatively different ones, forming, for example,
xyloglucan–cellulose bonds.
Following up the work on HvXET5, Fry et al. (2008a)
surveyed extracts from diverse land-plants and algae for
MXE activity. Curiously, one evolutionarily isolated genus,
Equisetum, gave extracts with very high MXE activity, often
exceeding its XET activity. Equisetum extracts acting on
MLG as donor showed several significant differences from
when acting on xyloglucan as donor, indicating that the
MXE-active protein is different from the major XET-active
XTHs. For example, (i) when XGOs were used as acceptor
substrates, different Equisetum extracts showed a consistent pattern of preferences among side chain-substituted
(1→4)-β-glucans (xyloglucan > water-soluble cellulose
acetate > HEC > CMC), whereas the rate with MLG as
Plant cell-wall enzymes | 3539
donor varied independently; (ii) MXE and XET activities
peaked in old and young Equisetum stems respectively; (iii)
MXE had a higher affinity for XXXGol (Km ~4 μM) than
any known XTH; and (iv) MXE and XET activities differed
in their oligosaccharide acceptor-substrate preferences,
for example XET activity 3-fold preferred XLLGol over
XXXGol, whereas MXE activity slightly preferred XXXGol
(Fry et al., 2008a).
Other possible homo- and hetero-transglycanases
The GHATAbase survey also indicated the presence in certain plant extracts (broad bean, pea, and cauliflower) of
trans-α-arabinanase activity, converting (1→5)-α-arabinooctaose (Ara8) to both smaller and larger products such as
Ara11–14 (Franková and Fry, 2011). The reaction may be of
the type
AAAA′ AAAA + AAAAAAAA ↔ AAAAAAAAAAAA + AAAA
where A = arabinose bond; ʹ = bond broken.
Other surveys of plant extracts returned negative data, e.g.
where the donor was xyloglucan and the prospective acceptors were galacto- or arabino-oligosaccharides that represent
RG-I side chains (Popper and Fry, 2008). Thus, no evidence
could be found for a hetero-transglycanase that might generate the xyloglucan–rhamnogalacturonan bonds detected
in muro (Thompson and Fry, 2000; Popper and Fry, 2005).
Negative data were also obtained with homogalacturonans as
donor and/or acceptor (García-Romera and Fry, 1994); this
can now be explained by the fact that all plant galacturonases
are inverting hydrolases (CAZy family GH28) and would
thus not readily evolve transglycanase capability.
An interesting high-throughput strategy for discovering transglycanases has been developed (Kosík et al., 2010).
Fourteen different polysaccharides (potential donors) were
printed onto a nitrocellulose surface, forming a ‘glycochip’,
which was then bathed in an Arabidopsis or Tropaeolum
enzyme extract containing 5 μM of an oligosaccharide–SR
(potential acceptor). After incubation, the chip was washed
in ethanol, which removed unreacted oligosaccharide, after
which any fluorescence on the chip was considered to represent a transglycanase product. With the Tropaeolum extract,
numerous donor:acceptor pairs led to apparent activity,
including some novel pairs, such as galactomannan:XGO,
glucuronoxylan:XGO,
galactomannan:MLGO,
and
xyloglucan:MLGO. Other pairs were also listed, for example
xyloglucan:cello-oligosaccharides, xyloglucan:laminari-oligosaccharides, and glucuronoxylan:galactomannan-oligosaccharides, but data in these cases are not shown. Conversely,
some substrate pairs do seem to reveal slight activity, especially at pH 7, for example arabinoxylan:XGO,
pectinate:XGO, arabinogalactan:XGO, and arabinan:XGO,
but these examples were not mentioned. Curiously, in the
arabinoxylan:XGO case, activity was only observed with arabinoxylan from wheat, not oat (supplementary figure S2 of
Kosík et al., 2010), raising the possibility that a contaminating polysaccharide in the wheat preparation was responsible.
Compared with Tropaeolum, Arabidopsis extracts gave far
fewer ‘hits’; for example, galactomannan:XGO activity was
undetectable. Boiled-enzyme controls verified the lack of
physical binding of the fluorescent oligosaccharides to the
‘printed’ polysaccharides. However, the study lacked a control testing whether polysaccharides co-extracted with the
enzymes (e.g. xyloglucan in the case of Tropaeolum seedlings)
might hydrogen-bond to some of the immobilized polysaccharides on the chip and give false-positives—fluorescent
spots that resemble novel hetero-transglycanase products but
are actually attributable to XET activity.
Stratilová et al. (2010) further showed that hetero-transglycanase activities of a purified pI-6.3 Tropaeolum XTH
represent side reactions due to relaxed substrate specificity,
the favoured activity being XET. Evidence for this conclusion was that low concentrations of added unlabelled XGOs
were sufficient to competitively interfere in hetero-reactions.
For example, the xyloglucan:MLGO–SR (MXE) reaction
was 93% inhibited by 1.9 mM unlabelled XGOs but only 35%
inhibited by a higher concentration (7.6 mM) of unlabelled
MLGOs.
Trans-α-xylosidase and other transglycosidase
activities
In principle, any retaining glycosidase can exhibit some transglycosidase activity at high substrate concentrations. For
example, 0.5–2.0 M substrates revealed trans-α-galactosidase
activity in a Prunus α-galactosidase (Dey, 1979), and 100 mM
nitrophenyl β-glucoside revealed trans-β-glucosidase activity in a barley β-glucosidase (Hrmova et al., 1998). These
are unphysiologically high concentrations. Transglycosidase
activities detectable at lower concentrations start to become
biologically significant. For example, a purified Tropaeolum
seed enzyme exhibited trans-β-glucosidase activity in a
TLC-based assay on a defined oligosaccharide, cellotetraose
[(1→4)-β-Glc4]. This activity transiently competed with concurrent β-glucosidase (exo-hydrolytic) activity when acting
on 5 mM cellotetraose, although hydrolysis strongly predominated at 2 mM, so it is uncertain whether the transglycosidase
activity would operate appreciably at physiological substrate
concentrations (Crombie et al., 1998).
In rice, both bglu1 and bglu2 (encoding β-glucosidases)
are highly expressed in young seedlings and mature nodes,
bglu1 also in flowers (Opassiri et al., 2003). Purified bglu1
hydrolysed both pNP-β-Glc and certain small oligosaccharides, optimally (1→3)-β-Glc2 and (1→4)-β-Glc4, and showed
moderate trans-β-glucosidase activity when these substrates
were present at 5 mM. This supports the theory that bglu1
can potentially generate new longer oligosaccharides from
shorter ones in vivo and this may play a role in recycling the
sugars released from the cell wall after germination or during
flower expansion (Opassiri et al., 2003). However, it is unclear
what biological benefit might accrue from resizing small
cello-oligosaccharides.
Incubation of 1.4 mM XXXG with diverse plant extracts
gave a series of products one, two, or three sugar residues larger
than the heptasaccharide starting material, indicating transglycosidase activity (Fig. 5). The virtual absence of free xylose
3540 | Franková and Fry
Fig. 5. TLC evidence for trans-α-xylosidase activity. The chromatogram shows products formed by the action of plant extracts on
1.4 mM XXXG (xyloglucan heptasaccharide; for further explanation of G, L, and X, see text). Pure substrate is shown in the left-hand
lane. Blue indicates smaller products remaining after transglycosylation plus any hydrolysis products; pink indicates products formed by
trans-α-xylosylation, during which xylose residue(s) are added to XXXG or to one of its major smaller products (pink arrows indicating
these trans-xylosylation reactions). DP, degree of polymerization (for example, DP10 indicates a decasaccharide); IP, isoprimeverose.
From Franková and Fry (2012a). Trans-[α]-xylosidase and trans-[β]-galactosidase activities, widespread in plants, modify and stabilise
xyloglucan structures. The Plant Journal, with permission of John Wiley & Sons.
among the reaction products indicates little α-xylosidase (exohydrolase) activity (Franková and Fry, 2012a). 3H-Labelled
substrates showed detectable transglycosylation at concentrations as low as 16 μM (in addition to hydrolysis, which is
favoured at such low substrate concentrations).
Definitive evidence for the trans-α-xylosidase reaction occurring was provided by dual-labelling experiments. With an aminolabelled (cationic) XGO as acceptor and [xylosyl-3H]XXXG as
donor, extracts of monocots (snowdrop, asparagus) and dicots
(chicory, parsley, cauliflower) all generated double-labelled (positively charged, radioactive) products (Franková and Fry, 2012a):
2012a). It is likely that polysaccharide-to-polysaccharide transfer
of single xylose residues can also occur. In the case of the XGO-toXGO reaction, the transferred α-xylose residue ends up attached to
an existing xylose residue (not to a glucose residue), forming a novel
xyloglucan trisaccharide unit, α-Xylp-(1→4)-α-Xylp-(1→6)-Glc,
assigned the sequence code-letter ‘V’ (Franková and Fry, 2012b).
It will be interesting to discover whether V also occurs in natural
xyloglucan.
A similar dual-labelling strategy using [Gal-3H]XXLG or
[Gal-3H]XLLG and XGO–NH2 gave evidence for trans-βgalactosidase activity in plant extracts. In contrast, [Fuc-3H]
With 37 μM [Xyl-3H]XXXG as donor plus 1 mM XGO–NH2 as
acceptor, transglycosylation exceeded xylosyl hydrolysis 1.6- to 7.3fold (lowest in cauliflower, highest in snowdrop), implying the presence of enzymes that favour transglycosylation. The extracts also
transferred α-xylose residues from [xylosyl-3H]XXXG to polysaccharide acceptor substrates (xyloglucan, water-soluble cellulose acetate, MLG, glucomannan, and arabinoxylan) (Franková and Fry,
XXFG gave no positively charged, radioactive products, indicating the absence of a trans-α-fucosidase activity – as predicted for CAZy class GH95 (inverting), which includes plant
α-1,2-fucosidases.
Panning for novel homo- and hetero-transglycosidase
activities is now feasible, given that simple but effective assays
are available.
Plant cell-wall enzymes | 3541
Phylogenetic variation
Although the genomic approach is valuable for revealing predicted proteins that may contribute to cell-wall restructuring, the strong possibility exists that additional hydrolase and
transglycosylase activities are being overlooked because their
genes are unidentified or incorrectly annotated. The discovery
of a unique endotransglucosylase activity (MXE) in Equisetum
(Fry et al., 2008a) emphasises the need to demonstrate biochemically the reactions catalysed by predicted wall enzymes.
Surveys of wall-acting enzyme activities thus usefully complement the ongoing description of the corresponding genes.
Among land plants, appreciable MXE activity is almost
confined to a single genus, Equisetum (the horsetails).
Equisetum probably qualifies as the most evolutionarily isolated genus of all living land-plants, its closest extant relatives having diverged >370 000 000 years ago (in the Upper
Devonian), at about the time when Tiktaalik, the almost-tetrapod fish, was groping its way into swamps. Real amphibians did not emerge until about 35 million years later, in the
Carboniferous. Equisetum has thus been evolving independently from all other plants for >370 My, so it is not surprising that it has acquired (or retained) some biochemical
peculiarities, including MXE.
The confinement of MXE to one genus, Equisetum, relates
to taxonomic specialization in cell-wall chemistry. Many of the
major discontinuities in plant evolution involved significant
alterations in wall polysaccharides (Popper, 2008; Fry, 2011;
Sørensen et al., 2011), which will have required new biosynthetic
genes but also driven the acquisition of novel enzymes involved
in wall restructuring. For example, MLG (one of the substrates
of MXE) is largely confined to the Poales, Equisetales, and
lichens (Trethewey et al., 2005; Fry et al., 2008b).4 Chemical
evidence for xyloglucan (e.g. its hydrolysis to yield isoprimeverose) is limited to embryophytes (not their sister group the
charophytes; Popper and Fry, 2003) (although immunolabelling
suggests that a chemically-related polysaccharide is present in
certain Chara cells, Domozych et al., 2009); low-fucose xyloglucan is characteristic of the Poales and the Solanales (Carpita
and Gibeaut, 1993; McDougall and Fry, 1994; Hoffman et al.,
2005); β-GalA-rich xyloglucans are widespread only in mosses
and liverworts (Peña et al., 2008), although they also occur in
angiosperm root-hairs (Peña et al., 2012); α-Arap in xyloglucan is known only from Equisetum and Selaginella (Peña et al.,
2008); and 3-O-methylrhamnose is a major wall component
only in charophytes, bryophytes, and homosporous lycopodiophytes (Popper et al., 2004), 3-O-methylgalactose being added
by the heterosporous lycopodiophytes (Popper et al., 2001).
These observations lead to the prediction that wall-modifying
enzymes should also vary between taxa.
This prediction was supported by an extensive study of
phylogenetic variation in wall polysaccharide-modifying
enzymes. The results are collated in GHATAbase (Franková
and Fry, 2011). First, great variation was noted in the occurrence of known enzyme activities between different plants,
4
in some cases correlating with taxonomic differences in
polysaccharide composition. Secondly, the results revealed
several new transglycanase and transglycosidase activities,
whose existence would not have been predicted by genomic
approaches (Mao et al., 2009) and whose biological roles
now invite exploration. In addition, GHATAbase (Table 1) is
a valuable resource for selecting plant tissues from which to
extract and study enzymes of interest.
Experimentally investigating wall enzyme
action in vivo
Genes predicted to encode wall-modifying enzymes may
or may not exhibit the predicted activity in vitro, and this
uncertainty can readily be explored by assays on the purified protein. But, another important question is whether the
enzyme actually catalyses any reaction (‘acts’) in vivo. Four
approaches to answering this puzzle, with different pros and
cons, are discussed in the following sections.
The mutation/RNAi approach
The first approach is to mutate (or incapacitate via RNAi) the
relevant gene and to observe whether any phenotypic change
ensues – which may indicate a direct or indirect role of the targeted gene. T-DNA insertion lines and RNAi plants provide useful plant material deficient in the protein (although, in the case
of multigene families, not necessarily the activity) of interest.
For example, knocking out AtXyl1, thought to be the only
Arabidopsis gene encoding α-xylosidase activity, resulted in (i)
a reduced ability of the seedlings to control the anisotropic
growth of several organs and (ii) xyloglucan with a reduced
proportion of Fuc and Gal, possibly affecting its interaction
with cellulose (Sampedro et al., 2010). This observation suggests that the AtXyl1 gene plays an important role in plant
growth. However, since a lack of α-xylosidase activity induced
a pleiotropic increase in other glycosidase activities (e.g.
β-galactosidase and α-fucosidase), it seems that gene silencing
does not reliably discriminate between direct and indirect roles
of xyloglucan-active exohydrolases (Sampedro et al., 2010).
Knocking out XTH31, a highly root-expressed gene that
encodes an XEH-active protein, caused a ~38% inhibition of
root elongation, suggesting that it acts in vivo and contributes to
root growth (Zhu et al., 2012). Comparison of an xth31 knockout mutant with the wild-type (Col-0) showed that normal
XTH31 expression has, surprisingly, several pleiotropic roles:
(i) increasing total extractable XET activity and in situ observable XET action, but not total extractable XEH activity; (ii)
promoting the elongation of roots by about 60%; (iii) increasing xyloglucan accumulation in the wall; and (iv) increasing the
Al3+-binding capacity of root cell walls. Interestingly, the 60%
element of growth attributable to XTH31 is strongly inhibited
by 5 μM Al3+, whereas the XTH31-independent element of
growth (that which continues unabated in the xth31 mutant) is
The occurrence of MLG in Selaginella is uncertain: immunological evidence supported it in S. moellendorffii (Harholt et al., 2012), but
chromatographic analysis suggested its absence in S. willdenowii (Xue and Fry, 2012).)
3542 | Franková and Fry
only slightly affected by 50 μM Al3+. These observations could
imply that XTH31, and/or some of its indirect effects, are specifically inhibited by Al3+.
XTH31 protein is detectable mainly in internal tissues of
the root’s elongation zone, whereas in-vivo XET action predominates in outer tissues (supplementary figures of Zhu
et al., 2012). This indicates that most XET action in roots
was not directly due to XTH31, in agreement with the observation that the heterologously produced protein has high
XEH and only very slight XET activity. In view of this, it
was surprising that xth31 mutant roots show diminished insitu XET action and low extractable XET activity. Knocking
out XTH31 indirectly diminishes the action and activity of
certain XET-active proteins such as XTH12 and XTH13
(Zhu et al., 2012). The findings emphasize that the mutation
approach, although demonstrating phenotypic consequences
of genes in vivo, does not guarantee that observed effects are
direct effects on restructuring of the expected substrate.
Clear evidence for a role of XET activity was also obtained
by the mutation approach in additional studies. XTH15 and
XTH17 are upregulated in Arabidopsis petioles in response
to dim light, an environmental stress that triggers rapid cell
elongation. Mutation of either of these genes abolished this
shade-avoidance response, suggesting a causal role, attributing
a biological function to two XET-active proteins (Sasidharan
et al., 2010). XTH18 is highly expressed in Arabidopsis roots, and
RNAi plants compromised in its expression exhibited a 16–18%
inhibition of root elongation rate and epidermal cell length
(Osato et al., 2006). This supports the idea that XTH18 contributes to root growth, suggesting important XET action in vivo.
An interesting study of AtBXL1 reveals some further
potential ambiguities in the genetic approach. This gene
encodes a β-xylosidase that is produced in vascular tissues,
and bxl1 antisense plants with decreased β-xylosidase activity
were found to have defective development (e.g. short siliquae
and curled leaf edges; Goujon et al., 2003). These observations might suggest that BXL1 exhibits β-xylosidase action in
vivo, contributing to normal plant growth. Antisense technology, however, is not guaranteed to target only a single gene,
and it remains possible that AtBXL2, which is 70% identical to AtBXL1, was co-suppressed. This suspicion was reinforced by the finding that bxl1-1, an insertional mutant, was
developmentally normal in its growth; bxl1-1 was, however,
defective in the swelling and release of seed-coat mucilage
and therefore in germination (Arsovski et al., 2009). Analysis
of the mutant’s mucilage showed 4.5-fold elevated proportions of 5-linked Araf residues in seed-coat RG-I and this,
coupled with the discovery that AtBXL1 is a bifunctional
β-d-xylopyranosidase/α-l-arabinofuranosidase (Minic et al.,
2004) led to the suggestion that its true action, at least in the
seed-coat, is α-arabinosidase. The bxl1-1 mutant seed-coat
mucilage showed no change in Xyl residues (Arsovski et al.,
2009). It remains to be seen whether the enzyme’s β-xylosidase
activity is manifested in β-xylosidase action in any other tissues in vivo. It is interesting that AtBXL1 appears to have
a role (seed-coat mucilage de-arabinosylation) not copied by
any of the other three (UniProtKB) putative α-arabinosidases
of Arabidopsis thaliana.
Tracing the fate of exogenous labelled model
substrates
A second, more direct, approach investigates enzyme action by
in-vivo observations on the behaviour of exogenous labelled oligomeric substrates which mimic wall polysaccharides but which
(unlike exogenous polysaccharides) can freely diffuse into contact with enzymes that are buried deep in the wall matrix.
Labelled XGOs, when fed to living plant tissues, act as acceptor substrates for XET action. The diagnostic product is a
labelled polysaccharide, easily distinguished from the oligosaccharide by its solubility properties. This product will be formed
only if an endogenous XET-active protein molecule can physically access an endogenous xyloglucan molecule (chemically
suitable as a donor substrate); the exogenous labelled XGO is
small enough to diffuse throughout the apoplast and come into
contact with the polysaccharide–enzyme complex. If the participating endogenous polysaccharide was hydrogen-bonded
near its non-reducing end to a microfibril, then the labelled
polysaccharide product will remain integrated within the cell
wall when washed in aqueous solutions. On the other hand,
if the donor polysaccharide was unattached to cellulose, or
hydrogen-bonded only near its reducing end, then the product
will be water-soluble – but will nevertheless remain in place if
the specimen is washed in 70% ethanol rather than water.
Several fluorescent and radioactive XGOs have been used in
this way to demonstrate XET action in vivo in various cell and
tissue systems. For example, 3H-labelled XGOs were shown to
become covalently linked to endogenous xyloglucan in cell cultures (Baydoun and Fry, 1989; Smith and Fry, 1991). Fluorescent
XGOs were applied similarly – with the advantage of ease of
localization but with less convenient quantification – in celery petioles (Fry, 1997), Arabidopsis root elongation zones (Vissenberg
et al., 2000), root hair initiation sites (Vissenberg et al., 2001),
specifically at the microfibril–matrix interface (Vissenberg et al.,
2005), in BY2 cultured tobacco cells (Ito and Nishitani, 1999),
in the developing G-layers of xylem (Nishikubo et al., 2007), in
opening carnation petals (Harada et al., 2011), and in cells of the
alga Chara (Van Sandt et al., 2007).
This approach is also useful to demonstrate the absence of
a postulated enzyme action in vivo. For instance, the suggestion that an XTH which exhibits slight MXE (hetero-transglycanase activity in vitro may endow barley tissues with MXE
action in vivo (Hrmova et al., 2007) was rejected on the basis
of experiments in which [3H]XGOs were fed to living barley
tissues. Action of endogenous XTH on endogenous barley
hemicelluloses generated [3H]xyloglucan (a high-Mr product
releasing [3H]XGOs when treated with xyloglucanase), confirming XET action, but did not form any [3H]MLG (which
would have released [3H]XGOs when treated with lichenase),
demonstrating the absence of appreciable MXE action in vivo
(Mohler et al., 2013). Identical experiments did demonstrate
substantial MXE action in Equisetum tissue.
This general approach is potentially capable of detecting
the in-vivo action of any of the other transglycanases whose
activities have been detected in vitro, for example trans-βxylanase (Franková and Fry, 2011; Johnston et al., 2013) and
trans-β-mannanase (Schröder et al., 2006).
Plant cell-wall enzymes | 3543
Feeding labelled oligosaccharides to living tissues gives
strong evidence that transglycanases act in vivo, but one
ambiguity remains: although action of endogenous enzyme
on endogenous donor polysaccharide is demonstrated, it
remains possible that no endogenous acceptor substrate
was accessible in vivo, in which case the transglycosylation reaction (begun by the formation of a polysaccharide–
enzyme complex) would not be consummated by transfer
of the polysaccharide to another wall component in vivo.
The only relevant part of the acceptor substrate is its nonreducing terminus, which for XET action should preferably
be an α-Xyl-(1→6)-Glc (isoprimeverose) unit, and it is possible that this terminus either is masked (inaccessible) within
the wall matrix (Marcus et al., 2010) or has been docked by
α-xylosidase action (Sampedro et al., 2010; Franková and
Fry, 2011; Günl and Pauly, 2011) or ‘end-capped’ by transα-xylosidase action (Franková and Fry, 2012a, b). Seemingly
minor modifications such as these, changing just the single
residue at the non-reducing extremity of a large polysaccharide chain, potentially take a whole xyloglucan molecule out
of play as far as its acceptor substrate role is concerned.
Nevertheless, this potential ambiguity in the interpretation
of experiments involving exogenous labelled XGOs does not
detract from their usefulness in providing evidence against
postulated enzyme actions, e.g. of MXE in barley (Mohler
et al., 2012) and of pectic transglycanase in cultured Rosa and
Acer cells (García-Romera and Fry, 1994).
Tracking the molecular ‘careers’ of endogenous
polysaccharides
A third approach by which to investigate whether wall
enzymes act in vivo is to track the molecular ‘careers’ of their
putative substrates (endogenous polysaccharides) and determine whether these undergo chemical changes of the predicted type. Such tracking is feasible after in-vivo feeding of
an exogenous, isotopically labelled, metabolic precursor that
labels a cohort of contemporaneously synthesized polysaccharide molecules, whose degradation/modification can then
be monitored in vivo at intervals thereafter.
Under these conditions, all three players in a transglycanase
reaction (enzyme, donor substrate, and acceptor) are endogenous. The method is particularly powerful if dual labelling
is applied. For example, plant cells are pregrown in substrate concentrations of a non-radioactive, density-labelled,
metabolically central precursor (e.g. glucose labelled with 2H
and/or 13C), so that all wall components become uniformly
‘heavy’. At time t, the tissue is transferred into medium containing ordinary glucose so that all future polysaccharides
synthesized are ‘light’. Soon after t, a tracer concentration
of radiolabelled precursor (e.g. [1-3H]Ara) is added. When
the resulting ‘hot, light’ polysaccharides reach the apoplast,
they may undergo interpolymeric transglycosylation with
the ‘cold, heavy’ molecules already there, yielding ‘hot, halfheavy’ products. Density of the 3H-polysaccharides is determined by isopycnic centrifugation in caesium trifluoroacetate.
By such experiments, and by altering the timing of the various medium shifts, clear evidence for XET action in vivo was
obtained (Thompson et al., 1997; Thompson and Fry, 2001).
The results supported XET action of two types: (i) ‘integrational transglycosylation’, in which part of a newly secreted
xyloglucan chain is grafted to a previously wall-bound one,
contributing to wall assembly; and (ii) ‘restructuring transglycosylation’, in which two xyloglucan molecules, both already
wall-bound, participate in the reaction, resulting in a remodelling of existing wall architecture, with probable wall loosening necessary for cell expansion.
Use of xenobiotics to inactivate a family of related wall
proteins
For a fourth approach, wall enzymes can be inactivated by exogenous inhibitory chemicals (xenobiotics), and the biological
consequences observed. In principle, this approach can provide
data comparable to those obtained from mutant or RNAi lines,
but with the advantage that a whole class of related wall enzyme
proteins may be targeted simultaneously. Since Arabidopsis has
33 XTH genes, attempts to produce a plant totally lacking XET
activity would require a line with mutations in all 33 genes.
Such a line would be tedious to generate and would possibly be
embryo lethal, precluding any insight into XET’s roles in postembryonic growth and development. However, a xenobiotic
targeting the active site common to all XTHs could be applied
at any desired stage of development, and the consequences of
a sudden block in XET action observed. For this reason the
Edinburgh Cell Wall Group is screening xenobiotic collections
on a high-throughput basis to search for inhibitors of XET and
other wall enzymes. The discovery of suitable xenobiotics will
not only provide a valuable tool for exploring the in-vivo roles of
wall enzymes, but, with wall targets being highly specific to the
plant kingdom, and in some cases to a narrow range of plant
taxa, they may also present novel opportunities for new selective herbicide development.
Concluding remarks
In conclusion, by focusing on plant cell-wall-localized glycosylhydrolases and the ability of some of these enzymes to catalyse transglycosylation reactions, this review draws attention
to interesting and largely overlooked activities likely to serve
important roles in the post-synthetic modification of plant cellwall polysaccharides in vivo. By considering the mechanism of
transglycosylation (versus hydrolysis), we highlight the fact that
the enzymic activities of some well-documented gene-products
may frequently be misinterpreted. An open mind is necessary
when studying transcript profiles and online databases to avoid
jumping to ill-founded conclusions about what reaction(s) a
wall enzyme may actually catalyse in vivo. To promote an evidence-based approach to elucidating their roles, this review has
summarized some of the methods currently available for tracing the behaviour of the enzymes’ substrates – mainly pectic
and hemicellulosic polysaccharides – in the walls of living plant
cells. It is predicted that application of such techniques will
open new insights into the mechanism and regulation of cell
expansion and other wall-centred aspects of plant physiology.
3544 | Franková and Fry
Acknowledgements
The authors thank the Leverhulme Foundation for supporting their work in this field.
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