Recent structural insights into the expanding world of carbohydrate-active enzymes

Article (PDF Available)inCurrent Opinion in Structural Biology 15(6):637-45 · January 2006with92 Reads
DOI: 10.1016/j.sbi.2005.10.008 · Source: PubMed
Abstract
Enzymes that catalyse the synthesis and breakdown of glycosidic bonds account for 1-3% of the proteins encoded by the genomes of most organisms. At the current rate, over 12 000 glycosyltransferase and glycoside hydrolase open reading frames will appear during 2006. Recent advances in the study of the structure and mechanism of these carbohydrate-active enzymes reveal that glycoside hydrolases continue to display a wide variety of scaffolds, whereas nucleotide-sugar-dependent glycosyltransferases tend to be grafted onto just two protein folds. The past two years have seen significant advances, including the discovery of a novel NAD+-dependent glycosidase mechanism, the dissection of the reaction coordinate of sialidases and a better understanding of the expanding roles of auxiliary carbohydrate-binding domains.
Recent structural insights into the expanding world of
carbohydrate-active enzymes
Gideon J Davies
1
, Tracey M Gloster
1
and Bernard Henrissat
2
Enzymes that catalyse the synthesis and breakdown of
glycosidic bonds account for 1–3% of the proteins encoded by
the genomes of most organisms. At the current rate, over
12 000 glycosyltransferase and glycoside hydrolase open
reading frames will appear during 2006. Recent advances in the
study of the structure and mechanism of these carbohydrate-
active enzymes reveal that glycoside hydrolases continue to
display a wide variety of scaffolds, whereas nucleotide-sugar-
dependent glycosyltransferases tend to be grafted onto just
two protein folds. The past two years have seen significant
advances, including the discovery of a novel NAD
+
-dependent
glycosidase mechanism, the dissection of the reaction
coordinate of sialidases and a better understanding of the
expanding roles of auxiliary carbohydrate-binding domains.
Addresses
1
Structural Biology Laboratory, Department of Chemistry, University of
York, Heslington, York YO10 5YW, UK
2
Architecture et Fonction des Macromole
´
cules Biologiques, UMR6098,
CNRS, Universite
´
s Aix-Marseille I & II, Case 932, 163 Avenue de Luminy,
13288 Marseille cedex 9, France
Corresponding author: Davies, Gideon J (davies@ysbl.york.ac.uk)
Current Opinion in Structural Biology 2005, 15:637–645
This review comes from a themed issue on
Catalysis and regulation
Edited by William N Hunter and Ylva Lindqvist
Available online 2nd November 2005
0959-440X/$ see front matter
# 2005 Elsevier Ltd. All rights reserved.
DOI 10.1016/j.sbi.2005.10.008
Introduction
Carbohydrates in the form of glycoproteins, glycolipids
and polysaccharides play fundamental roles in the cell
physiology and development of microbes, plants and
animals. The enzymes that cleave and build the glyco-
sidic bonds of glycoconjugates, oligosaccharides and poly-
saccharides comprise a group of enzymes that act on the
most structurally diverse substrates in nature. These
‘carbohydrate-active enzymes’ (CAZymes) have been
classified into several families based on amino acid
sequence similarity [1]. This classification system, which
presently counts over 200 families of glycosidases, gly-
cosyltransferases, polysaccharide lyases and carbohydrate
esterases (Figure 1), is available from the continuously
updated carbohydrate-active enzyme database (CAZy) at
http://afmb.cnrs-mrs.fr/CAZY/. A feature that makes
these families particularly useful is that, although accom-
modating varying substrate specificities, they correlate
with several structure-based properties of the enzymes,
such as an essentially conserved fold and active site
geometry. With the advent of genome sequencing, such
a classification system has great predictive power.
Here, we review the emerging genomic data and show
how glycosidases continue to be a rapidly advancing field,
with many new structures emerging each year that,
viewed in light of chemical data, shed new insight into
reaction mechanism. Glycosyltransferase research pro-
ceeds at a slower pace, whilst the emerging field of sugar
deacetylation brings new life to old mechanisms. Aux-
iliary carbohydrate-binding domains find more applica-
tions, reflecting their diverse substrate specificities in
vitro and, increasingly, in vivo.
A promenade through genomic data
The sequence-based classification of carbohydrate-active
enzymes provides an efficient tool for the competent
annotation (e.g. prediction of the general function, fold
and mechanism) of open reading frames (ORFs) found
during genome sequencing. Examination of the comple-
ment of carbohydrate-active enzymes within the different
(non-archaeal) genomes listed in the CAZy database
reveals that, for most organisms, 1–3% of their genes
encode glycoside hydrolases or glycosyltransferases
(Figure 2). There is massive gene loss accompanying
the acquisition of parasitic or symbiotic lifestyles [2];
indeed, some organisms, such as Ehrlichia ruminantium,
Francisella tularensis and Guillardia theta, appear to have
lost all their carbohydrate-active enzymes (at least all
those related to known ones). At the other end of the
spectrum, some organisms clearly have more carbohy-
drate-active enzyme encoding genes than average. The
champion, by total number, is certainly Arabidopsis thali-
ana; however, even it will soon be dwarfed by the poplar
genome. In terms of percentage of the genome devoted to
carbohydrate-active enzymes, the human gastrointestinal
tract bacteria Bacteroides thetaiotaomicron (6.6%), Bacter-
oides fragilis NCTC 9343 (4.8%) and Bifidobacterium
longum (3.6%) reign supreme, hinting at our intimate
dependence on these organisms and their enzymes. It
is clear that both conventional and genome sequencing
are providing a wealth of ORFs for enzymes acting in the
synthesis and degradation of carbohydrates. Not surpris-
ingly, both structural and mechanistic work lags some-
what behind. The CAZy classification does at least
provide a framework upon which to discuss recent
advances in our understanding of the structure and
www.sciencedirect.com Current Opinion in Structural Biology 2005, 15:637–645
638 Catalysis and regulation
Figure 1
The world of carbohydrate-active enzymes. Many classes of enzymes are active in the formation, modification and breakdown of glycosides.
For the purposes of this review, we consider glycoside hydrolases, which are responsible for the hydrolysis of the glycosidic bond, and whose
action can result in net retention or inversion of the configuration of the anomeric carbon; glycosyltransferases, which use the energy derived
from activated sugar donors to drive glycosidic bond synthesis (typically the activating group is a nucleotide or lipid-phosphate); carbohydrate
esterases/deacetylases, which perform the de-O and de-N acetylation of acetylated sugars; and polysaccharide lyases (not discussed in this
review), which catalyse the b-elimination reaction on uronic acid glycosides.
Figure 2
Rapidly expanding genome sequence information provides a wealth of carbohydrate-active enzyme sequences. (a) As of September 2005, over
280 organisms have had their genome fully sequenced, with the sequences of over 600 genomes currently under preparation (Genome Online
Database; http://www.genomesonline.org/). Excluding archaea, organisms typically use 1–3% of their genome for carbohydrate synthesis and
degradation, with the synthetic enzymes showing a much better correlation with genome size (R
2
= 0.7) than the hydrolases (R
2
= 0.48).
Plants and human intestinal bacteria tend to lie above the line, with parasites such as Plasmodium (and Homo sapiens) below. (b) The number
of sequences of these enzymes becoming available is expanding rapidly (glycoside hydrolases, closed circles; glycosyltransferases, open circles);
at the current rate, over 12 000 glycosyltransferase and glycoside hydrolase ORFs will appear in 2006.
Current Opinion in Structural Biology 2005, 15 :637–645 www.sciencedirect.com
mechanism of enzymes responsible for the synthesis and
degradation of carbohydrates.
Glycoside hydrolases: a marketplace of
different structures
Glycoside hydrolases (see Figure 1) are, arguably, the
best-characterised enzymes active on disaccharides, oli-
gosaccharides and polysaccharides. The CAZy classifica-
tion lists 100 sequence-derived families and there are
three-dimensional structural representatives for over 60
of these! These enzymes continue to show a vast array of
tertiary scaffolds, as we originally observed over 10 years
ago [3]. It is beyond the scope of this review to comment
on all the recently determined three-dimensional struc-
tures. Notable achievements have included structure
determinations for members of long-sought-after enzyme
classes, including fucosidase [4], invertase [5], levansu-
crase [6], agarase [7], dextranase [8] and the GH31
a-glycosidases [9]. In the plant cell-wall context, two
particularly insightful works come from studies of xylo-
glucan-metabolising enzymes. The structure of the GH74
xyloglucanase [10] provides a first glimpse of the con-
sortium of enzymes involved in plant xyloglucan degra-
dation, whereas one of the most significant structures of
recent years, that of the xyloglucan endotransferase XET,
provides the first structural view of how plants manipulate
and remodel xyloglucan during plant growth and expan-
sion [11

]. The XET work also lays the platform for the
exciting manipulation of xyloglucan in the production of
‘designer’ paper products [12
].
Ballet within active sites: new mechanistic
insights into glycosidic bond hydrolysis
Perhaps the most exciting applications of three-dimen-
sional structure occur when the crystal structure acts in
tandem with solution work to define new mechanistic
paradigms. The two standard mechanisms of glycosidic
bond cleavage, leading to either inversion or retention of
the configuration of the anomeric carbon, were originally
outlined by Koshland in 1953 [13]. With only a few subtle
variations, these proposals have stood the test of time, and
have received widespread investigation and review [14–
16]. Recently, a fundamentally different glycosidase
mechanism has been unveiled through structure deter-
mination and kinetic analyses of the NAD
+
and divalent
metal ion dependent GH4 glycosidases [17–19]. Three-
dimensional structures revealed that the NAD
+
moiety
sat just below C3 of the sugar, with a tyrosine residue
above C2 (Figure 3). Kinetic isotope effect measurements
[20

] demonstrated that abstraction of a hydride from C3
and a proton from C2 were both partially rate limiting. An
overall mechanism can thus be proposed in which hydride
abstraction at C3 generates a ketone, which has the effect
of acidifying the C2 proton, allowing deprotonation by the
tyrosine residue accompanied by acid-catalysed elimina-
tion of the glycosidic oxygen and formation of a 1,2-
unsaturated intermediate. This a,b-unsaturated species
then undergoes base-catalysed attack by water to gener-
ate a 3-keto glucose derivative, which is then reduced by
the ‘on-board’ NADH, returning the enzyme to its active
form and completing the reaction cycle. This work on
GH4 enzymes is one of few recent examples in which
structure and mechanistic study have combined to deliver
an unexpected result; the other significant example is the
dissection of the mechanism of action of sialidases, work
that may have ramifications for us all.
The media interest in the likely ‘H5N1’ influenza pan-
demic has probably not escaped anyone’s attention. It is
unlikely, however, that many have realised that ‘H’ and
‘N’ refer to a carbohydrate-binding protein and a carbo-
hydrate-active enzyme, respectively. H is a haemagglu-
tinin, which binds cell-surface sialic acid; haemagglutinin
mutations are responsible for influenza crossing the spe-
cies divide (elegantly described at the three-dimensional
level in the context of the 1918 influenza by Gamblin et al.
[21

]). N is a neuraminidase, or ‘sialidase’, which hydro-
lyse the glycosidic bond between sialic acid and other
sugars, allowing viral release. Given the implicit involve-
ment of sialidases, not only in influenza but also in other
serious disease processes, it is encouraging that there have
been major advances in the structural and mechanistic
biology of these enzymes in recent times. Sialidases were
always a mechanistic conundrum; they act with retention
of anomeric configuration, but do not possess an aspartate
or glutamate residue appropriately positioned to act as a
nucleophile in a double-displacement mechanism.
Recent outstanding work, harnessing both the trapping
methodology of Withers and co-workers [22
] and the
brilliant structure determination of a trans-sialidase by
Alzari and co-workers [23

], has shown that, for this
system at least, the sialidase mechanism involves the
formation and subsequent breakdown of a tyrosyl-
enzyme intermediate (Figure 4)[24
]. Given the impor-
tance of mechanistic inhibitors of sialidases, examples in
clinical use include Relenza and Tamiflu, such structural
and mechanistic studies of these enzymes cannot be
overstated. The sialidase story will run and run. Bennet
and colleagues [25] have subsequently shown that some
mutants of the nucleophilic tyrosine both remain active
and still perform catalysis despite the absence of a
nucleophile for the double displacement. Similarly, struc-
ture determination of the very unusual polysialic-acid-
degrading bacteriophage K1F endosialidase reveals a
much-compromised active centre grafted onto a triple-
helical stalk [26](Figure 4); however, the enzyme is still
functional — again demonstrating that sialidases have not
yet given up all their secrets.
Glycosyltransferases: a slow-moving cart
Glycosyltransferases catalyse the transfer of an activated
donor sugar to an appropriate acceptor, typically another
sugar, lipid, protein or small molecule (Figure 1). The
CAZy classification describes 78 sequence-based families
Carbohydrate-active enzymes Davies, Gloster and Henrissat 639
www.sciencedirect.com Current Opinion in Structural Biology 2005, 15:637–645
of glycosyltransferases [27], comprising 15 800 ORFs. It
is immediately noticeable that, in contrast to the glyco-
side hydrolase work, the structural biology of glycosyl-
transferases is a slow-moving vehicle, with just 20 of the
78 families having a three-dimensional structural repre-
sentative to date (in contrast to the success of structural
genomics with other classes of sugar-active enzymes,
high-throughput approaches have yielded just a single
glycosyltransferase).
Historically, following on from Vrielink and Freemont’s
pioneeringworkonDNAb-glucosyltransferase (GT-63)
back in 1994 [28], it was a full five years before Charnock
and Davies [29] provided a new structural example of
these activated-sugar-dependent enzymes. Indeed,
since 1999, only a handful of glycosyltransferase struc-
tures have been reported, with the following families
having representative three-dimensional structures
(only post-2003 examples are cited): GT-1 [30
], GT-
2, GT-5 [31], GT-6 [32], GT-7, GT-8, GT-9, GT-13,
GT-15 [33], GT-20 [34], GT-27 [35

], GT-28 [36],
GT-35, GT-42 [37

], GT-43, GT-44 [38], GT-63,
GT-64 [39

], GT-72 [40]andGT-78[41
]. Thus far,
all these structures have revealed just two canonical
folds, termed GT-A and GT-B, after their initial obser-
vation in the SpsA and DNA b-glucosyltransferase struc-
tures, respectively (Figure 5).
At the three-dimensional level, the past two years have
seen several considerable achievements in glycosyltrans-
ferase structural biology. The structure of the first sialyl-
transferase (from family GH-42) revealed an unusual
variant of the GT-A fold [37

]. A particularly pleasing
structure is that of the GT-27 UDP-GalNAc:polypeptide-
N-acetylgalactosaminyltransferase-T1, which is involved
in mucin biosynthesis [35

]. This structure solution of an
extremely important enzyme in glycobiology also faced
the difficulties posed by a dynamic twin-domain archi-
tecture in which the glycosyltransferase domain is
appended to a carbohydrate-binding module. The solu-
tion of the mannosylglycerate synthase structure by Flint
et al. is informative not merely as a ligand-complexed
form of a GT-A fold transferase that acts with retention of
anomeric configuration, but also more as an example of
how modern high-throughput activity screens can begin
to shed light upon the catalytic function of glycosyltrans-
640 Catalysis and regulation
Figure 3
An exciting new mechanism of glycosidic bond cleavage by family GH4 glycosidases. These enzymes harness NAD
+
as a transient redox
catalyst, oxidising C3 to a ketone and thereby acidifying C2 of the glycone moiety. Sequential 2,1 elimination of the aglycone and 1,2 addition of
water, followed by re-reduction of the ketone yield the product sugar, with net retention of anomeric configuration. Such a mechanistic formulation
can only be revealed through the synergistic interplay of three-dimensional structure [17–19] and chemical [20

] work.
Current Opinion in Structural Biology 2005, 15 :637–645 www.sciencedirect.com
ferases [41
]. Genes encoding uncharacterised glycosyl-
transferases keep on accumulating in huge numbers. One
of the major challenges facing glycobiologists in this field,
especially those working on bacterial and plant enzymes,
is to divine the donor and acceptor specificities from a
myriad of possibilities, based on only a tiny proportion of
biochemically characterised cases.
Carbohydrate esterases: mechanisms from
the catacombs?
The CAZy classification lists 14 different families of
carbohydrate esterases/deacetylases. Biologically, these
enzymes are involved in the removal of O- (ester) and
N-acetyl moieties from carbohydrates; indeed, sugar dea-
cetylases display similar catalytic strategies to those
Carbohydrate-active enzymes Davies, Gloster and Henrissat 641
Figure 4
The developing area of sialidase structure and mechanism. The reaction mechanism of sialidases, which act with net retention of anomeric
configuration, has always been a controversial area, not least as there was no suitably positioned aspartate or glutamate to act as a nucleophile
in a double-displacement mechanism. (a) Recent exciting work by the Withers [22
] and Alzari [24
] groups has trapped a catalytically competent
tyrosyl-enzyme intermediate of the trypanosomal trans-sialidase, showing that this is the likely intermediate during sialidase catalysis. (b) The
unusual bacteriophage K1F endosialidase possesses three ‘sialidase’ domains grafted onto a triple-helical stalk [26], yet the active centre of
these domains is compromised, showing that sialidases still have many secrets to give up. Study of sialidase structure and mechanism is
essential if we are to build on pioneering drugs such as (c) Tamiflu and Relenza, which may find increasing use in any forthcoming influenza
pandemic.
Figure 5
The two glycosyltransferase folds identified thus far. (a) GT-A fold, as exemplified by the GT-27 UDP-GalNAc:polypeptide-N-
acetylgalactosaminyltransferase-T1, which is involved in mucin biosynthesis [35

]. This outstanding structure determination had to overcome the
challenges associated with a flexible multimodular protein with an appended CBM (right). (b) GT-B fold, as exemplified by the recently determined
structure of GtfD, which is involved in the synthesis of the antibiotic vancomycin [30
]. Some families, such as the sialyltransferases [37

] of family GH-
42, have unusual variants of these folds. The structures are colour ramped from N terminus (blue) to C terminus (red), with any ligands present shown in
ball-and-stick representation.
www.sciencedirect.com Current Opinion in Structural Biology 2005, 15:637–645
employed by more classical esterases and peptidases. In
fact, the similarity of different sugar and non-sugar
esterases, and the promiscuity of their substrate specifi-
city make the carbohydrate esterase classification in many
cases less insightful and predictive than the hydrolase,
transferase and lyase families. The majority of carbohy-
drate esterases/deacetylases whose structures have been
reported display a classical b/a/b ‘serine protease’ fold, as
revealed by three-dimensional structures of the enzymes
from family CE-1 (bacterial ferulate esterases), CE-5
(acetyl xylan esterases), CE-7 (multifunctional and xyloo-
ligosaccharide deacetylases [42]), the plethora of enzymes
from family CE-10, the mycolyltransferase ‘antigen 85C’
and the non-classified fungal ferulate esterases [43]. A
small deviation from this canonical fold is displayed by
the CE-12 rhamnogalacturonan acetylesterase [44]. Pec-
tin methylesterases from family CE-8 use a different
mechanism, in which a twin-aspartate catalytic centre
is grafted onto a right-handed parallel b helix [45]. These
enzymes may also be considered slightly unusual in that it
is the sugar that forms the acid, rather than the ‘R’ group
(see Figure 1).
Some sugar deacetylase structures have also revealed
both single and double metal ion catalytic centres (well
reviewed in a general context in [46]). Notable examples
include the LpxC zinc-dependent UDP-3-O-acetyl-N-
acetylglucosamine deacetylases from family CE-11,
which present a classical zinc hydrolase site on a novel
a/b framework [47,48]; the single-zinc CE-14 N-acetyl-1-
D-myo-inosityl-2-amino-2-deoxy-a-D-glucopyranoside
deacetylase [49]; the family CE-4 deacetylases [50],
sometimes refereed to as ‘NodB homologs’, whose mem-
bers are involved in the deacetylation, amongst other
things, of peptidoglycan, chitin, rhizobial Nod factors
and xylan; and the twin-metal, urease-like CE-9 N-acet-
ylglucosamine-6-phosphate deacetylase [51].
Carbohydrate-binding domains: arguing
about a role?
One of the features of enzymes active in the degradation
of oligosaccharides and polysaccharides, occasionally also
reflected in the synthetic enzymes, is the presence of one
or more non-catalytic modules involved in the targeting of
these biocatalysts to their cognate substrates. These
domains, termed ‘carbohydrate-binding modules’ or
CBMs, potentiate the activity of the parent enzyme
against insoluble substrates. Currently, the CAZy classi-
fication lists 43 families of characterised CBMs (reviewed
in a structural context in [52
]), but this is probably a gross
under-estimate as there are dozens of families of domains
of unknown function that will prove to be CBMs when
their biochemical properties have been interrogated.
Given the recent review, we limit ourselves to a discus-
sion of only two recent aspects of CBM work that are of
particular importance.
There has long been controversy over whether the CBM
itself has ‘disruptive’ but non-hydrolytic activity on the
substrate, thus promoting catalysis by increasing access to
the target carbohydrate, in addition to aiding adsorption.
Recently, however, it has been shown, convincingly, that
a (non-enzyme appended) chitin-binding CBM domain,
CBP21 from Serratia marcescens, promotes hydrolysis of
crystalline chitin via non-hydrolytic degradation of the
substrate [53
]. A similarly disputed feature of CBMs is
why prokaryotes have evolved many CBMs with appar-
ently identical targets. We are only now beginning to
appreciate subtle differences in the substrate specificity
of these domains, with work showing the binding of
642 Catalysis and regulation
Figure 6
From understanding to exploiting CBMs. Recent years have seen a vast explosion in our knowledge of enzyme-appended CBMs, with many
diverse specificities reported and an array of structures often displaying different sugar-binding modes (reviewed in [52
]). The Gilbert and Knox
groups have demonstrated that different CBMs, with apparently similar specificities in vitro, actually target different cellular structures in vivo.The
figure shows indirect immunofluorescence microscopy of (a) CBM2b1-2, (b) CBM15 and (c) CBM35 binding to transverse sections of tobacco
stem (adapted from [54

]), showing cortical parenchyma (cp) and pith parenchyma (pp), in addition to vascular tissues (e, epidermis; p, phloem
fibres; x, xylem vessels). Arrow pairs indicate files of developing xylem vessel elements (CBM2b1-2 binds to these). CBM35 binds to files of
parenchyma cells between them. The large arrow head indicates cells associated with internal phloem. Scale bar = 100 mm.
Current Opinion in Structural Biology 2005, 15 :637–645 www.sciencedirect.com
apparently similar domains to very different regions of
plant cell walls in vivo [54

](Figure 6). Such work, which
will no doubt expand greatly, begins to shed light not only
on the raison d’e
ˆ
tre of CBMs but also on how they may be
exploited in cellular mapping processes.
Opening the great gates to future challenges
In the ten years since we first reviewed structural aspects
of carbohydrate-active enzymes [3], much has occurred,
but what challenges remain? Evidently, there is a major
effort to provide structural representatives for the remain-
ing glycoside hydrolase families; will they reveal yet more
novel three-dimensional folds converging on just a hand-
ful of different catalytic strategies? In higher organisms,
we can expect to see the massive expansion of ‘chemical
genetics’ approaches to the identification [55

,56], dis-
section and subsequent inhibition [57

] of important
hydrolytic enzymes. For glycosyltransferases, major chal-
lenges exist at all levels from protein expression through
to functional annotation. Compared to glycoside hydro-
lases [58], our detailed knowledge of glycosyltransferase
mechanism is extremely poor and, if these enzymes are
ever to become serious targets for therapy, much work
remains. Indeed, we have no structural insight into lipid-
phosphate-dependent glycosyltransferases at all and yet
these are some of the most significant enzymes on earth.
Sharon [59] described glycobiology as ‘‘the last frontier of
molecular and cell biology’’. This sentiment remains true
on many levels, not least our three-dimensional insight
into the processes of glycosidic bond formation.
Acknowledgements
The authors would like to thank all those who made suggestions on this
article and apologise to those whose work could not be accommodated.
GJD is a Royal Society University Research Fellow.
References and recommended reading
Papers of particular interest, published within the annual period of
review, have been highlighted as:
of special interest
 of outstanding interest
1. Coutinho PM, Henrissat B: Carbohydrate-active enzymes: an
integrated approach.In Recent Advances in Carbohydrate
Engineering. Edited by Gilbert HJ, Davies GJ, Svensson B,
Henrissat B. Royal Society of Chemistry; 1999:3-12.
2. Henrissat B, Deleury E, Coutinho PM: Glycogen metabolism
loss: a common marker of parasitic behaviour in bacteria?
Trends Genet 2002, 18 :437-440.
3. Davies G, Henrissat B: Structures and mechanisms of glycosyl
hydrolases. Structure 1995, 3:853-859.
4. Sulzenbacher G, Bignon C, Nishimura T, Tarling CA, Withers SG,
Henrissat B, Bourne Y: Crystal structure of Thermotoga
maritima alpha-
L-fucosidase - insights into the catalytic
mechanism and the molecular basis for fucosidosis. J Biol
Chem 2004, 279:13119-13128.
5. Alberto F, Bignon C, Sulzenbacher G, Henrissat B, Czjzek M:
The three-dimensional structure of invertase (beta-
fructosidase) from Thermotoga maritima reveals a bimodular
arrangement and an evolutionary relationship between
retaining and inverting glycosidases. J Biol Chem 2004,
279:18903-18910.
6. Meng G, Futterer K: Structural framework of fructosyl transfer
in Bacillus subtilis levansucrase. Nat Struct Biol 2003,
10:935-941.
7. Allouch J, Helbert W, Henrissat B, Czjzek M: Parallel
substrate binding sites in a beta-agarase suggest a novel
mode of action on double-helical agarose. Structure 2004,
12:623-632.
8. Larsson AM, Andersson R, Stahlberg J, Kenne L, Jones TA:
Dextranase from Penicillum minioluteum: reaction course,
crystal structure, and product complex. Structure 2003,
11:1111-1121.
9. Lovering AL, Lee SS, Kim YW, Withers SG, Strynadka NCJ:
Mechanistic and structural analysis of a family 31 alpha-
glycosidase and its glycosyl-enzyme intermediate . J Biol Chem
2005, 280:2105-2115.
10. Yaoi K, Kondo H, Noro N, Suzuki M, Tsuda S, Mitsuishi Y: Tandem
repeat of a seven-bladed beta-propeller domain in
oligoxyloglucan reducing-end-specific cellobiohydrolase.
Structure 2004, 12:1209-1217.
11.

Johansson P, Brumer H, Baumann MJ, Kallas AM, Henriksson H,
Denman SE, Teeri TT, Jones TA: Crystal structures of a poplar
xyloglucan endotransglycosylase reveal details of
transglycosylation acceptor binding. Plant Cell 2004,
16:874-886.
Arguably one of the most sought-after three-dimensional structures in
plant biochemistry. XET catalyses the cleavage and remodelling of plant
cell-wall xyloglucan. This masterful work reveals the three-dimensional
structure and the basis of its acceptor substrate specificity.
12.
Brumer H, Zhou Q, Baumann MJ, Carlsson K, Teeri TT: Activation
of crystalline cellulose surfaces through the chemoenzymatic
modification of xyloglucan. J Am Chem Soc 2004,
126:5715-5721.
Building upon the structural work described in [11

], the XET enzyme is
used to graft functional groups onto cellulosic (paper) surfaces without
damaging the integrity of the cellulose.
13. Koshland DE: Stereochemistry and the mechanism of
enzymatic reactions. Biol Rev 1953, 28:416-436.
14. Zechel DL, Withers SG: Glycosidase mechanisms: anatomy of a
finely tuned catalyst. Acc Chem Res 2000, 33:11-18.
15. Vocadlo DJ, Davies GJ, Laine R, Withers SG: Catalysis by hen
egg-white lysozyme proceeds via a covalent intermediate.
Nature 2001, 412:835-838.
16. Vasella A, Davies G, Bo
¨
hm M: Glyco sidase mechanisms.
Curr Opin Chem Biol 2002, 6:619-629.
17. Lodge JA, Maier T, Liebl W, Hoffmann V, Strater N: Crystal
structure of Thermotoga maritima alpha-glucosidase AglA
defines a new clan of NAD
+
-dependent glycosidases.
J Biol Chem 2003, 278:19151-19158.
18. Rajan SS, Yang X, Collart F, Yip VLY, Withers SG, Varrot A,
Thompson J, Davies GJ, Anderson WF: NAD-dependent
hydrolysis by family 4 glycosidases involves a novel
elimination mechanism. Structure 2004, 12:1619-1629.
19. Varrot A, Yip VLY, Li Y, Rajan SS, Yang X, Anderson W,
Thompson J, Withers SG, Davies GJ: NAD
+
and metal-ion
dependent hydrolysis by family 4 glycosidases: structural
insight into specificity for phospho-b-
D-glucosides. J Mol Biol
2005, 346:423-435.
20.

Yip VLY, Varrot A, Davies GJ, Rajan SS, Yang X, Thompson J,
Anderson WF, Withers SG: An unusual mechanism of glycoside
hydrolysis involving redox and elimination-steps by a family 4
b-glycosidase from Thermotoga maritima. J Am Chem Soc
2004, 126:8354-8355.
This work highlights the essential synergy between ‘physical organic’
enzymology and three-dimensional structure. Kinetic isotope effect mea-
surements at the C2 and C3 positions with solvent isotope exchange
allow postulation of a novel reaction mechanism for these NAD
+
-depen-
dent glycosidases, whose structures are described in [17–19].
21.

Gamblin SJ, Haire LF, Russell RJ, Stevens DJ, Xiao B, Ha Y,
Vasisht N, Steinhauer DA, Daniels RS, Elliot A et al.: The structure
and receptor binding properties of the 1918 influenza
hemagglutinin. Science 2004, 303:1838-1842.
Carbohydrate-active enzymes Davies, Gloster and Henrissat 643
www.sciencedirect.com Current Opinion in Structural Biology 2005, 15:637–645
This article, and another in the same issue, analyses the three-dimen-
sional structures of influenza virus haemagglutinins from various ‘flu
strains, including the fabled 1918 ’flu. Mutations in the ligand-binding
site allow ’flu to cross the species barrier by changing the specificity from
sialic acid linked a-2,3 to galactose in avian intestines to the a-2,6 linkage
of human respiratory tract glycans.
22.
Watts AG, Damager I, Amaya ML, Buschiazzo A, Alzari P,
Frasch AC, Withers SG: Trypanosoma cruzi trans-sialidase
operates through a covalent sialyl-enzyme intermediate:
tyrosine is the catalytic nucleophile. J Am Chem Soc 2003,
125:7532-7533.
The first experimental demonstration that sialidases operate through the
formation and subsequent breakdown of a covalent tyrosyl-enzyme
intermediate. It will be interesting to see if this is demonstrated for a
range of different sialidases.
23.

Buschiazzo A, Amaya MF, Cremona ML, Frasch AC, Alzari PM:
The crystal structure and mode of action of trans-sialidase, a
key enzyme in Trypanosoma cruzi pathoge nesis. Mol Cell 2002,
10:757-768.
A tour-de-force structure determination of a medically important enzyme,
brought about through insightful biochemistry and the construction of
surface ‘crystallisation mutants. Sialic acid is shown to trigger a con-
formational change that increases affinity for the acceptor and aids
transglycosylation, which is crucial to trypanosomal infection.
24.
Amaya MF, Watts AG, Damager T, Wehenkel A, Nguyen T,
Buschiazzo A, Paris G, Frasch AC, Withers SG, Alzari PM:
Structural insights into the catalytic mechanism of
Trypanosoma cruzi trans-sialidase. Structure 2004,
12:775-784.
An elegant and revealing series of structural snapshots along the reaction
coordinate of trans-sialidase, including the trapping of the covalent
tyrosyl-enzyme intermediate.
25. Watson JN, Newstead S, Narine A, Taylor G, Bennet AJ: Two
nucleophilic mutants of the Micomonospora viridifaciens
sialidase operate with retention of configuration by two
different mechanisms. ChemBioChem 2005, in press.
26. Stummeyer K, Dickmanns A, Muhlenhoff M, Gerardy-Schahn R,
Ficner R: Crystal structure of the polysialic acid-degrading
endosialidase of bacteriophage K1F. Nat Struct Mol Biol 2005,
12:90-96.
27. Coutinho P, Deleury E, Davies GJ, Henrissat B: An evolving
hierarchical family classification for glycosyltransferases.
J Mol Biol 2003, 328:307-317.
28. Vrielink A, Ru
¨
ger W, Driessen HPC, Freemont PS: Crystal
structure of the DNA modifying enzyme b-glucosyltransferase
in the presence and absence of the substrate uridine
diphosphoglucose. EMBO J 1994, 13:3413-3422.
29. Charnock SJ, Davies GJ: Structure of the nucleotide-
diphospho-sugar transferase, SpsA from Bacillus subtilis,in
native and nucleotide-complexed forms. Biochemistry 1999,
38:6380-6385.
30.
Mulichak AM, Lu W, Losey HC, Walsh CT, Garavito RM:
Crystal structure of vancosaminyltransferase GtfD from
the vancomycin biosynthetic pathway: interactions with
acceptor and nucleotide ligands. Biochemistry 2004,
43:5170-5180.
One in a series of perceptive papers from this team, who have provided
unparalleled views of the modification of antibiotics by glycosylation. The
GtfD structure was determined in complex with both nucleotide and the
desvancosaminyl vancomycin acceptor, giving an in-depth understand-
ing of specificity and catalysis.
31. Buschiazzo A, Ugalde JE, Guerin ME, Shepard W, Ugalde RA,
Alzari PM: Crystal structure of glycogen synthase: homologous
enzymes catalyze glycogen synthesis and degradation.
EMBO J 2004, 23:3196-3205.
32. Lee HJ, Barry CH, Borisova SN, Seto NOL, Zheng RXB,
Blancher A, Evans SV, Palcic MM: Structural basis for the
inactivity of human blood group O-2 glycosyltransferase.
J Biol Chem 2005, 280:525-529.
33. Lobsanov YD, Romero PA, Sleno B, Yu BM, Yip P, Herscovics A,
Howell PL: Structure of Kre2p/Mnt1p - a yeast alpha 1,2-
mannosyltransferase involved in mannoprotein biosynthesis.
J Biol Chem 2004, 279:17921-17931.
34. Gibson R, Tarling CA, Roberts S, Withers SG, Davies GJ: The
donor subsite of trehalose-6-phosphate synthase: binary
complexes with UDP-glucose and UDP-2-deoxy-2-fluoro
glucose at 2A
˚
resolution. J Biol Chem 2004, 279:1950-1955.
35.

Fritz TA, Hurley JH, Trinh LB, Shiloach J, Tabak LA: The
beginnings of mucin biosynthesis: the crystal structure of
UDP-GalNAc: polypeptide alpha-N-
acetylgalactosaminyltransferase-T1. Proc Natl Acad Sci USA
2004, 101:15307-15312.
The authors report the structure of an extremely important enzyme, one
that displays a modular architecture featuring a CBM appended to the
glycosyltransferase domain. The structure confirms early suggestions
that this family of enzymes, which act with retention of anomeric con-
figuration during catalysis, would be similar in three-dimensional struc-
ture to GT-2 inverting enzymes, as proposed in [29].
36. Hu Y, Chen L, Ha S, Gross B, Falcone B, Walker D, Mokhtarzadeh
M, Walker S: Crystal structure of the MurG: UDP-GlcNAc
complex reveals common structural principles of a
superfamily of glycosyltransferases. Proc Natl Acad Sci USA
2003, 100:845-849.
37.

Chiu CPC, Watts AG, Lairson LL, Gilbert M, Lim D, Wakarchuk
WW, Withers SG, Strynadka NCJ: Structural analysis of the
sialyltransferase CstII from Campylobacter jejuni in complex
with a substrate analog. Nat Struct Mol Biol 2004, 11:163-170.
Cell-surface sialic acid is one of the most important glycosylations in
nature. This paper reports the first sialyltransferase structure determina-
tion and was consequently one of the American Chemical Society
‘Chemical Highlights’ of 2004. The enzyme transfers sialic acid to cell-
surface glycoproteins and glycolipids. It has an unusual deviation from
the canonical GT-A fold, with the donor interactions revealed through
crystallisation with a non-transferrable DMP-sialic acid mimic.
38. Reinert DJ, Jank T, Aktories K, Schulz GE: Structural basis for the
function of Clostridium difficile toxin B. J Mol Biol 2005,
351:973-981.
39.

Pedersen LC, Dong J, Taniguchi F, Kitagawa H, Krahn JM,
Pedersen LG, Sugahara K, Negishi M: Crystal structure of an
alpha 1,4-N- acetylhexosaminyltransferase (EXTL2), a
member of the exost osin gene family involved in heparan
sulfate biosynthesis. J Biol Chem 2003, 278:14420-14428.
A fine structure determination and very interesting piece of scientific
writing. All those working on retaining glycosyltransferases should read
this paper, which reports the first structures of members of the exostosin
gene family and provides an important structural dissection of the
mechanisms of heparan synthesis.
40. Larivie
`
re L, Sommer N, More
´
ra S: Structural evidence of a
passive base-flipping mechanism for AGT, an unusual GT-B
glycosyltransferase. J Mol Biol 2005, 352:139-150.
41.
Flint J, Taylor E, Yang M, Bolam DN, Tailford LE, Martinez-Flietes
C, Dodson EJ, Davis BG, Gilbert HJ, Davies GJ: Structural
dissection and high-throughput screening of
mannosylglycerate synthase. Nat Struct Mol Biol 2005,
12:608-614.
This work suggests one possible route to the functional dissection of
glycosyltransferase activity. Three-dimensional structure is mute without
solution characterisation of activity. Nowhere is this more true than in
glycosyltransferase studies, not least because the two known folds are
also adopted by enzymes that are not glycosyltransferases.
42. Vincent F, Charnock SJ, Verschueren KHG, Turkenburg JP,
Scott DJ, Offen WA, Roberts S, Pell G, Gilbert HJ, Davies GJ et al.:
Multifunctional xylooligosaccharide/cephalosporin C
deacetylase revealed by the hexameric structure of the
Bacillus subtilis enzyme at 1.9A
˚
resolution. J Mol Biol 2003,
330:593-606.
43. Hermoso JA, Sanz-Aparicio J, Molina R, Juge N, Gonzalez R,
Faulds CB: The crystal structure of feruloyl esterase a from
Aspergillus niger suggests evolutive functional convergence
in feruloyl esterase family. J Mol Biol 2004, 338:495-506.
44. Mølgaard A, Kauppinen S, Larsen S: Rhamnogalacturonan
acetylesterase elucidates the structure and function of a new
family of hydrolases. Structure 2000, 8:373-383.
45. Jenkins J, Mayans O, Smith D, Worboys K, Pickersgill RW: Three-
dimensional structure of Erwinia chrysanthemi pectin methyl
esterase reveals a novel esterase active site. J Mol Biol 2001,
305:951-960.
644 Catalysis and regulation
Current Opinion in Structural Biology 2005, 15 :637–645 www.sciencedirect.com
46. Hernick M, Fierke CA: Zinc hydrolases: the mechanisms of
zinc-dependent deacetylases. Arch Biochem Biophys 2005,
433:71-84.
47. Whittington DA, Rusche KM, Shin H, Fierke CA, Chistianson DW:
Crystal structure of LpxC, a zinc-dependent deacetylase
essential for endotoxin biosynthesis. Proc Natl Acad Sci USA
2003, 100:8146-8150.
48. Coggins BE, Li X, McClerren AL, Hindsgaul O, Raetz CRH, Zhou P:
Structure of the lpxC deacetylase with a bound substrate
analog inhibitor. Nat Struct Biol 2003, 10:645-651.
49. Maynes JT, Garen C, Cherney MM, Newton G, Arad D, Av-Gay Y,
Fahey RC, James MNG: The crystal structure of 1-
D-myo-
inosityl-2-acetamido-2-deoxy-alpha-
D-glucopyranoside
deacetylase (MshB) from Mycobacterium tuberculosis reveals
a zinc hydrolase with a lactate dehydrogenase fold. J Biol
Chem 2003, 278:47166-47170.
50. Blair DE, Schuttelkopf AW, MacRae JA, van Aalten DMF:
Structure and metal-dependent mechanism of peptidoglycan
deacetylase, a Streptococcal virulence factor. Proc Natl Acad
Sci USA 2005, in press.
51. Vincent F, Yates D, Garman E, Davies GJ, Brannigan JA: The 3-D
structure of the N-acetylglucosamine-6-phosphate
deacetylase, NagA, from Bacillus subtilis: a member of the
urease superfamily. J Biol Chem 2004, 279:2809-2816.
52.
Boraston AB, Bolam DN, Gilbert HJ, Davies GJ: Carbohydrate-
binding modules: fine tuning polysaccharide recognition.
Biochem J 2004, 382:769-781.
A long-overdue review of CBMs, their three-dimensional structures and
their mechanism of binding.
53.
Vaaje-Kolstad G, Houston DR, Riemen AHK, Eijsink VGH, van
Aalten DMF: Crystal structure and binding properties of the
Serratia marcescens chitin-binding protein CBP21. J Biol
Chem 2005, 280:11313-11319.
This elegant work clearly demonstrates a highly significant enhancement
of enzyme activity when the CBM is added in trans’. Furthermore,
electron micrographs show clear physical impact on the substrate by
the chitin-binding module. A rare piece of unambiguous work in this
controversial area.
54.

McCartney L, Gilbert HJ, Bolam DN, Boraston AB, Knox JP:
Glycoside hydrolase carbohydrate-binding modules as
molecular probes for the analysis of plant cell wall polymers.
Anal Biochem 2004, 326:49-54.
The fact that many prokaryotes harness very different families of CBMs on
their enzymes has long been confusing. This work not only reveals that
different CBM families target different ‘substructures’ of plant cell-wall
polysaccharides, but also provides a method to exploit this specificity for
the analysis of cellular ‘glyco-architecture’.
55.

Vocadlo DJ, Hang HC, Kim EJ, Hanover JA, Bertozzi CR:
A chemical approach for identifying O-GlcNAc-modified
proteins in cells. Proc Natl Acad Sci USA 2003,
100:9116-9121.
An elegant ‘chemical biology’ approach to analysing the massively
important O-GlcNAc modification in eukaryotes. One can expect to
see similar approaches harnessed to study many of the complex glyco-
sylation reactions in the cell.
56. Vocadlo DJ, Bertozzi CR: A strategy for functional proteomic
analysis of glycosidase activity from cell lysates. Angew Chem
Int Ed Engl 2004, 43:5338-5342.
57.

Macauley MS, Whitworth GE, Debowski AW, Chin D, Vocadlo DJ:
O-GlcNAcase uses substrate-assisted catalysis - kinetic
analysis and development of highly selective mechanism-
inspired inhibitors. J Biol Chem 2005, 280:25313-25322.
Again a vision of the future using the O-GlcNAc modification as an
example. A panel of inhibitors both define the reaction mechanism and
allow specific intervention.
58. Davies GJ, Ducros VM-A, Varrot A, Zechel DL: Mapping the
conformational itinerary of b-glycosidases by X-ray
crystallography. Biochem Soc Trans 2003, 31:523-527.
59. Sharon N: The conquest of the last frontier of molecular and
cell biology. Biochimie 2001, 83:555.
Carbohydrate-active enzymes Davies, Gloster and Henrissat 645
www.sciencedirect.com Current Opinion in Structural Biology 2005, 15:637–645
    • "Carbohydrates are important natural molecules presented in their free moieties or in association in glycoproteins, glycolipids, and polysaccharides, playing fundamental roles in the cell physiology and development of all organisms [1]. The enzymes that cleave or, inversely, mediate the ligation of glycosidic bonds of glycoconjugates, oligosaccharides, and polysaccharides can be classified by two different systems. "
    [Show abstract] [Hide abstract] ABSTRACT: Glycoside hydrolases (GH) are enzymes capable to hydrolyze the glycosidic bond between two carbohydrates or even between a carbohydrate and a non-carbohydrate moiety. Because of the increasing interest for industrial applications of these enzymes, the immobilization of GH has become an important development in order to improve its activity, stability, as well as the possibility of its reuse in batch reactions and in continuous processes. In this review, we focus on the broad aspects of immobilization of enzymes from the specific GH families. A brief introduction on methods of enzyme immobilization is presented, discussing some advantages and drawbacks of this technology. We then review the state of the art of enzyme immobilization of families GH1, GH13, and GH70, with special attention on the enzymes β-glucosidase, α-amylase, cyclodextrin glycosyltransferase, and dextransucrase. In each case, the immobilization protocols are evaluated considering their positive and negative aspects. Finally, the perspectives on new immobilization methods are briefly presented.
    Full-text · Article · Aug 2016
    • "Glycosyltransferases catalyze the transfer of sugar moieties from activated donor molecules to specific acceptor molecules such as sugars, lipids, proteins, or small molecules including phenylpropanoids. Information on GHs and GTs, as well as polysaccharide lyases and carbohydrate esterases, can be found in the CarbohydrateActive enZymes (CAZy) database (Davies et al., 2005) and a specific PlantCAZyme database is now also available (Ekstrom et al., 2014). CAZy takes into account similarities in amino acid sequences, 3D-structures, sugar donors, transferred sugars, acceptors, catalytic mechanisms (inverting or retaining mechanisms) and more recently the combination of modules which can be catalytic or not (Lombard et al., 2014). "
    [Show abstract] [Hide abstract] ABSTRACT: The phenylpropanoid pathway in plants is responsible for the biosynthesis of a huge amount of secondary metabolites derived from phenylalanine and tyrosine. Both flavonoids and lignins are synthesized at the end of this very diverse metabolic pathway, as well as many intermediate molecules whose precise biological functions remain largely unknown. The diversity of these molecules can be further increased under the action of UDP-glycosyltransferases (UGTs) leading to the production of glycosylated hydroxycinnamates and related aldehydes, alcohols and esters. Glycosylation can change phenylpropanoid solubility, stability and toxic potential, as well as influencing compartmentalization and biological activity. (De)-glycosylation therefore represents an extremely important regulation point in phenylpropanoid homeostasis. In this article we review recent knowledge on the enzymes involved in regulating phenylpropanoid glycosylation status and availability in different subcellular compartments. We also examine the potential link between monolignol glycosylation and lignification by exploring co-expression of lignin biosynthesis genes and phenolic (de)glycosylation genes. Of the different biological roles linked with their particular chemical properties, phenylpropanoids are often correlated with the plant's stress management strategies that are also regulated by glycosylation. UGTs can for instance influence the resistance of plants during infection by microorganisms and be involved in the mechanisms related to environmental changes. The impact of flavonoid glycosylation on the colour of flowers, leaves, seeds and fruits will also be discussed. Altogether this paper underlies the fact that glycosylation and deglycosylation are powerful mechanisms allowing plants to regulate phenylpropanoid localisation, availability and biological activity
    Full-text · Article · May 2016
    • "For the GT-A fold enzymes the N-and C-terminal domains show dissimilar architecture. The N-terminal domain consists of several β-sheets, which all are flanked by α-helical Rossmann folds and are is responsible for recognition of the sugar-nucleotide donor (Davies et al., 2005; Jank et al., 2007; Mittler et al., 2007; Erb et al., 2009). The C-terminal domain mainly contains mixed β-sheets and is responsible for the binding of the acceptor molecule. "
    [Show abstract] [Hide abstract] ABSTRACT: The enzyme subclass of glycosyltransferases (GTs; EC 2.4) currently comprises 97 families as specified by CAZy classification. One of their important roles is in the biosynthesis of disaccharides, oligosaccharides, and polysaccharides by catalyzing the transfer of sugar moieties from activated donor molecules to other sugar molecules. In addition GTs also catalyze the transfer of sugar moieties onto aglycons, which is of great relevance for the synthesis of many high value natural products. Bacterial GTs show a higher sequence similarity in comparison to mammalian ones. Even when most GTs are poorly explored, state of the art technologies, such as protein engineering, domain swapping or computational analysis strongly enhance our understanding and utilization of these very promising classes of proteins. This perspective article will focus on bacterial GTs, especially on classification, screening and engineering strategies to alter substrate specificity. The future development in these fields as well as obstacles and challenges will be highlighted and discussed.
    Full-text · Article · Feb 2016
Show more