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

Recent structural insights into the expanding world of carbohydrate-active enzymes Gideon J Davies1, Tracey M Gloster1 and Bernard Henrissat2 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 carbohydrateactive enzymes reveal that glycoside hydrolases continue to display a wide variety of scaffolds, whereas nucleotide-sugardependent 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 ([email protected])

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 glycosidic bonds of glycoconjugates, oligosaccharides and polysaccharides 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, glycosyltransferases, 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 www.sciencedirect.com

these families particularly useful is that, although accommodating 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 proceeds at a slower pace, whilst the emerging field of sugar deacetylation brings new life to old mechanisms. Auxiliary carbohydrate-binding domains find more applications, 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 complement 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 carbohydrate-active enzyme encoding genes than average. The champion, by total number, is certainly Arabidopsis thaliana; 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%), Bacteroides 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 surprisingly, both structural and mechanistic work lags somewhat behind. The CAZy classification does at least provide a framework upon which to discuss recent advances in our understanding of the structure and Current Opinion in Structural Biology 2005, 15:637–645

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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 (R2 = 0.7) than the hydrolases (R2 = 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

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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, oligosaccharides and polysaccharides. The CAZy classification 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 structures. Notable achievements have included structure determinations for members of long-sought-after enzyme classes, including fucosidase [4], invertase [5], levansucrase [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 xyloglucan-metabolising enzymes. The structure of the GH74 xyloglucanase [10] provides a first glimpse of the consortium of enzymes involved in plant xyloglucan degradation, 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 expansion [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-dimensional 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 determination and kinetic analyses of the NAD+ and divalent metal ion dependent GH4 glycosidases [17–19]. Threedimensional 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 elimination of the glycosidic oxygen and formation of a 1,2unsaturated intermediate. This a,b-unsaturated species www.sciencedirect.com

then undergoes base-catalysed attack by water to generate 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 pandemic 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 carbohydrate-active enzyme, respectively. H is a haemagglutinin, which binds cell-surface sialic acid; haemagglutinin mutations are responsible for influenza crossing the species 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 hydrolyse the glycosidic bond between sialic acid and other sugars, allowing viral release. Given the implicit involvement 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 tyrosylenzyme intermediate (Figure 4) [24]. Given the importance 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, structure determination of the very unusual polysialic-aciddegrading bacteriophage K1F endosialidase reveals a much-compromised active centre grafted onto a triplehelical 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 Current Opinion in Structural Biology 2005, 15:637–645

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.

of glycosyltransferases [27], comprising 15 800 ORFs. It is immediately noticeable that, in contrast to the glycoside hydrolase work, the structural biology of glycosyltransferases is a slow-moving vehicle, with just 20 of the 78 families having a three-dimensional structural representative 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 pioneering work on DNA b-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 structures have been reported, with the following families having representative three-dimensional structures (only post-2003 examples are cited): GT-1 [30], GT2, 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] and GT-78 [41]. Thus far, Current Opinion in Structural Biology 2005, 15:637–645

all these structures have revealed just two canonical folds, termed GT-A and GT-B, after their initial observation in the SpsA and DNA b-glucosyltransferase structures, respectively (Figure 5). At the three-dimensional level, the past two years have seen several considerable achievements in glycosyltransferase structural biology. The structure of the first sialyltransferase (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:polypeptideN-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 architecture in which the glycosyltransferase domain is appended to a carbohydrate-binding module. The solution 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 glycosyltranswww.sciencedirect.com

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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.

ferases [41]. Genes encoding uncharacterised glycosyltransferases 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 deacetylases display similar catalytic strategies to those

Figure 5

The two glycosyltransferase folds identified thus far. (a) GT-A fold, as exemplified by the GT-27 UDP-GalNAc:polypeptide-Nacetylgalactosaminyltransferase-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 GH42, 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

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642 Catalysis and regulation

employed by more classical esterases and peptidases. In fact, the similarity of different sugar and non-sugar esterases, and the promiscuity of their substrate specificity make the carbohydrate esterase classification in many cases less insightful and predictive than the hydrolase, transferase and lyase families. The majority of carbohydrate 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 xylooligosaccharide 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]. Pectin 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-Nacetylglucosamine 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-1D -myo-inosityl-2-amino-2-deoxy-a- D-glucopyranoside deacetylase [49]; the family CE-4 deacetylases [50], sometimes refereed to as ‘NodB homologs’, whose members 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-acetylglucosamine-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 classification 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 discussion 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 apparently identical targets. We are only now beginning to appreciate subtle differences in the substrate specificity of these domains, with work showing the binding of

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

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Carbohydrate-active enzymes Davies, Gloster and Henrissat 643

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’eˆtre of CBMs but also on how they may be exploited in cellular mapping processes.

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.

Opening the great gates to future challenges

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 alphaglycosidase and its glycosyl-enzyme intermediate. J Biol Chem 2005, 280:2105-2115.

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 remaining glycoside hydrolase families; will they reveal yet more novel three-dimensional folds converging on just a handful of different catalytic strategies? In higher organisms, we can expect to see the massive expansion of ‘chemical genetics’ approaches to the identification [55,56], dissection and subsequent inhibition [57] of important hydrolytic enzymes. For glycosyltransferases, major challenges exist at all levels from protein expression through to functional annotation. Compared to glycoside hydrolases [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 lipidphosphate-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.

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644 Catalysis and regulation

This article, and another in the same issue, analyses the three-dimensional 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 pathogenesis. 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 conformational 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 nucleotidediphospho-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 understanding 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 1,2mannosyltransferase involved in mannoprotein biosynthesis. J Biol Chem 2004, 279:17921-17931. Current Opinion in Structural Biology 2005, 15:637–645

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-Nacetylgalactosaminyltransferase-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 configuration during catalysis, would be similar in three-dimensional structure 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 determination and was consequently one of the American Chemical Society ‘Chemical Highlights’ of 2004. The enzyme transfers sialic acid to cellsurface 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 exostosin 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: Threedimensional structure of Erwinia chrysanthemi pectin methyl esterase reveals a novel esterase active site. J Mol Biol 2001, 305:951-960. www.sciencedirect.com

Carbohydrate-active enzymes Davies, Gloster and Henrissat 645

46. Hernick M, Fierke CA: Zinc hydrolases: the mechanisms of zinc-dependent deacetylases. Arch Biochem Biophys 2005, 433:71-84.

electron micrographs show clear physical impact on the substrate by the chitin-binding module. A rare piece of unambiguous work in this controversial area.

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.

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’.

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-myoinosityl-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,

www.sciencedirect.com

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 glycosylation 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 mechanisminspired 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.

Current Opinion in Structural Biology 2005, 15:637–645