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ScienceDirect A sweet new wave: structures and mechanisms of enzymes that digest polysaccharides from marine algae Jan-Hendrik Hehemann1, Alisdair B Boraston2 and Mirjam Czjzek3,4 Marine algae contribute approximately half of the global primary production. The large amounts of polysaccharides synthesized by these algae are degraded and consumed by microbes that utilize carbohydrate-active enzymes (CAZymes), thus creating one of the largest and most dynamic components of the Earth’s carbon cycle. Over the last decade, structural and functional characterizations of marine CAZymes have revealed a diverse set of scaffolds and mechanisms that are used to degrade agars, carrageenan, alginate and ulvanpolysaccharides from red, brown and green seaweeds, respectively. The analysis of these CAZymes is not only expanding our understanding of their functions but is enabling the enhanced annotation of (meta)-genomic data sets, thus promoting an improved understanding of microbes that drive this marine component of the carbon cycle. Furthermore, this information is setting a foundation that will enable marine algae to be harnessed as a novel resource for biorefineries. In this review, we cover the most recent structural and functional analyses of marine CAZymes that are specialized in the digestion of macro-algal polysaccharides. Addresses 1 Massachusetts Institute of Technology, Department of Civil and Environmental Engineering, 15 Vassar Street, Bldg 48-108, Cambridge, MA 02139, USA 2 Biochemistry and Microbiology, University of Victoria, PO Box 3055 STN CSC, Victoria, BC, Canada V8W 3P6 3 Sorbonne Universite´s, UPMC, Universite´ Paris 06, France 4 CNRS UMR 8227, Integrative Biology of Marine Models, Station Biologique de Roscoff, CS 90074, F-29688 Roscoff cedex, Bretagne, France Corresponding authors: Hehemann, Jan-Hendrik ([email protected]), Czjzek, Mirjam ([email protected])
Current Opinion in Structural Biology 2014, 28:77–86 This review comes from a themed issue on Carbohydrate–protein interactions and glycosylation Edited by Harry Brumer and Harry J Gilbert
http://dx.doi.org/10.1016/j.sbi.2014.07.009 0959-440X/# 2014 Elsevier Ltd. All rights reserved.
Introduction About one-half of the global net primary production, which amounts to an estimated 104.9 pentagrams of www.sciencedirect.com
carbon per year, is generated by marine algae [1]. Marine heterotrophic microbes digest the produced organic matter, such as high molecular weight algal polysaccharides, into low molecular weight substrates. This is amongst the largest and fastest catabolic biotransformations on earth and is key to the carbon cycle by returning this photosynthetically fixed carbon back to the atmosphere. Indeed, the ocean has been compared to ‘a global digester, where almost all organic matter injected from surface waters (0–100 m) is respired to inorganic nutrients’ [2] and only a ‘small’ fraction escapes rapid microbial consumption. That this phenomenon exists and is critical to both ocean health and earth climate regulation is well appreciated; however, the microbial processes governing this turnover are not well understood, particularly at the molecular level. For example, on average, marine genomic or metagenomic data sets contain genes encoding up to 20% more hypothetical proteins with unknown functions and structures, or ‘orphans’, than their terrestrial counterparts [3]. Countless marine microorganisms associated with algae contain unexplored enzymes, such as novel glycoside hydrolases (GH) and polysaccharide lyases (PL), that participate in important metabolic pathways and whose characterization holds the potential to provide new insights into marine carbon recycling. The CAZy database (www.cazy.org) groups carbohydrateactive enzymes (CAZymes) that have a common ancestor into families of proteins and presently includes 133 GH families and 33 PL families [4,5]. Recent years have seen increasing interest in the analysis of CAZymes that degrade marine polysaccharides, mainly those from seaweeds. The first structure determined for such an enzyme was the kappa-carrageenase from the marine bacterium Pseudoalteromonas carrageenovora, which was isolated for its capacity to grow on carrageenan-gels [6]. While this enzyme belongs to the large, multi-specific family GH16, in which both ‘terrestrial’ and ‘marine’ enzymes occur [7,6,8,9,10], many more recently identified marine polysaccharide specific CAZymes have laid the basis for several novel and marine-dominant GH and PL families, such as b-agarases in GH50, GH86, and GH118; i-carrageenase in GH82; aagarases in GH96 and GH117; and alginate lyases in PL7, PL15, and PL17. As the database of CAZymes becomes increasingly populated with known and putative enzymes involved in the breakdown of marine algal polysaccharides some of the important questions directing the examination of these Current Opinion in Structural Biology 2014, 28:77–86
78 Carbohydrate–protein interactions and glycosylation
Table 1 CAZy famillies containing enzymes active on marine polysaccharides and their respective PDB accession codes CAZy family GH16
Protein
GH50
k-Carrageenase (Pseudoaltermonas carragenovora) b-Agarase A (Zobellia galactanivorans) b-Agarase B (Zobellia galactanivorans) b-Agarase D (Zobellia galactanivorans) b-Porphyranase (Bacteroides plebeius) b-Porphyranase A (Zobellia galactanivorans) b-Porphyranase B (Zobellia galactanivorans) b-1,3-Glucanase (Mycobacterium marinum M) Laminarinase (Thermotoga maritima) Laminarinase R (Rhodothermus marinus) Laminarinase A (Pyrococcus furiosus) Laminarinase A (Zobellia galactanivorans) Exo-b-agarase D (Saccharophagus degradans)
GH82
i-Carrageenase (Alteromonas fortis)
GH86 GH105 GH117
b-Porphyranase (Bacteroides plebeius) Unsaturated b-glucuronyl hydrolase (Nonlabens ulvanivorans) a-1,3-L-(3,6-Anhydro)-galactosidase (Zobellia galactanivorans) a-1,3-L-Neoagarobiose hydrolase (Saccharophagus degradans) a-1,3-L-(3,6-Anhydro)-galactosidase (Bacteroides plebeius) Endo-guluronate lyase (Zobellia galactanivorans) Exo-alginate lyase (Zobellia galactanivorans) Exo-alginate lyase (Agrobacterium fabrum) Oligoalginate lyase (Saccharophagus degradans)
PL7 PL15 PL17
PDB codes 1DYP 1O4Y; 1URX a 1O4Z; 4ATF 4ASM 4AWD 4ATE; 3ILF 3JUU 4PQ9 3AZX; 3AZZ 3ILN 2VY0 4BQ1; 4BOW 4BQ2; 4BQ3 4BQ4; 4BQ5 1H80; 1KTW 3LMW 4AW7 4CE7 3P2N 3R4Y; 3R4Z 4AK5; 4AK7 3ZPY 4BE3 3A0O; 3AFL 4NEI; 4OJZ
A complete overview of the structural state of all CAZyme families so far can be found in Lombard et al. [5]. For all GH16 enzymes, only one additional pdb code of a substrate-complex is given, when existent.
a
enzymes are: What are the precise specificities of these enzymes? What is the molecular basis behind enzymatic recognition of the unique structures of marine polysaccharides? What is the biochemistry underlying the enzymatic cleavage of glycosidic bonds in marine polysaccharides? The answers to these questions are helping us to understand how microbes process the billions of tons of polysaccharides that are produced annually by algae in the sea and informing how this process may change in future marine systems [11]. Importantly, such research is also providing biotechnological tools to produce tailored oligosaccharides with therapeutic potential and generating the biocatalysts required for efficient use of algal biomass as feedstocks for biorefinement processes [12]. Here we provide an update on the most recent advances in understanding enzymes involved in marine polysaccharide degradation (Table 1).
Seaweed degrading CAZymes for biorefinery applications In the quest for alternative biofuels seaweeds are a promising compliment to, or even substitute for, land plant biomass as a feedstock. Algae grow faster and do not impinge on arable land. Furthermore, seaweeds build their cell walls with higher amounts of gel forming polysaccharides that are highly hydrated and, relative to common ligno-cellulosic biomass, much easier to digest with enzymes. For example, depending on the type of seaweed, the cell walls of these macro-algae (Figure 1) Current Opinion in Structural Biology 2014, 28:77–86
contain up to 40% agarose, porphyran, carrageenan, alginate, or ulvan polysaccharides, most of which form hydrogels [13,14]. Recently, this advantage was successfully exploited by transferring an alginate-specific metabolic locus from Vibrio splendidus 12B01 [15] into a bioengineered Escherichia coli strain, resulting in a bacterium able to convert alginates from brown seaweeds into high yields of bioethanol [16]. This example illustrates the great potential of marine polysaccharide specific CAZymes for bioenergy production. Optimization of this approach to convert algal alginate into biofuel, however, and extension of the concept to utilize seaweeds rich in other polysaccharides requires ongoing structural and functional characterization of algae degrading GHs and PLs from marine microbes.
Hydrolytic enzyme systems for the digestion of red seaweeds The matrix polysaccharides of red algae are either agars (in agarophytes) or carrageenan (in carragenophytes). Enzymes degrading the latter polysaccharide were discovered early on during CAZyme prospecting and are extensively reviewed elsewhere [17]. Although bagarases have been characterized for some time, their structural analysis is a relatively recent development, as is the discovery and characterization of a-agarases. Agars are galactans comprising L-galactose that is a-1,3 linked with D-galactose (G). In agarose, the L-galactose is www.sciencedirect.com
Structures and mechanisms of marine CAZymes Hehemann, Boraston and Czjzek 79
Figure 1
2 1
3
4
3. GH117
1. GH16 OH HO
O
2. GH50
O
O
agarose
O
OH
O
O OH OHOH
GH16, GH86
OH
OH O HO
O O
porphyran
OH OH
O
O
O
OH
HO OH
O
O
OH
O O
OH
O
O
O
O
–O3S
OH
O –O3S
O
OH
1. PL7 OH
–OOC O
O OH
–OOC O HO
O OH
O
2. PL15, PL17, PL7 AlyA5 OH O
HO O
O OH
alginate
O COO-
–OOC OH
–OOC
GH105
–OOC O HO OH
H 3C O
OH
O
ulvan O
–O3S
O O
O HO
H3C O
O O
–O3S
OH
OH Current Opinion in Structural Biology
Red, green and brown algae contain different types of polysaccharides that are degraded by dedicated enzymatic systems. Left: Coast of rocky beach near Roscoff (Brittany, France) at low tide shows different types of seaweeds. Most visible are brown seaweeds of Fucus spp. and sugar kelp Saccharina lattissima Right: Close-up of left showing a patch with diversity of seaweeds present in this ecosystem. Inset legend: (1,2) the red seaweeds Porphyra spp. and Chondrus spp. respectively; (3) the green seaweed Ulva spp.; (4) the brown seaweed Fucus spp. The major cell wall matrix polysaccharides are shown. Agarose and porphyran are abundant in red seaweeds, alginate is present in brown seaweeds and ulvan comes from green seaweeds. These polysaccharides are degraded by glycoside hydrolases and polysaccharide lyases belonging to different families as indicated above the glycosidic bonds. The sequence in which these enzymes operate is indicated by numbers. For example, in alginate degradation the PL15, PL17 and PL7AlyA5 remove unsaturated monosaccharides from the non-reducing end of oligosaccharides produced by endo-acting Pl7 alginate lyases that initiate digestion.
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80 Carbohydrate–protein interactions and glycosylation
3,6-anhydro-L-galactose (LA) while in porphyran it is Lgalactose-6-sulfate (L6S). The resulting neoagarobiose and neoporphyrobiose disaccharides are connected by b-1,4 glycosidic linkages in agarose and porphyran, respectively (Figure 1) [18]. To degrade these polysaccharides marine microbes use a variety of enzymes to specifically hydrolyze the b-glycosidic and a-glycosidic linkages. Those that have been structurally characterized are covered in more detail below and discussed within the context of agarolytic systems.
Agarose degradation At present, b-agarases belong to families GH16, GH50, GH86, and GH118, while and a-agarases belong to families GH96 and GH117. Those that are structurally characterized are in GH16, GH50, and GH117. b-Agarases in GH16. The first three-dimensional structures of two b-agarases were of the enzymes ZgAgaA and ZgAgaB, which originate from the marine bacterium Zobellia galactanivorans. These enzymes belong to family GH16, have a b-jelly-roll-fold, and catalyze hydrolysis with a catalytic mechanism that retains the stereochemistry at the anomeric carbon and utilizes two catalytic glutamate residues [19–21]. Their endo-mode of action, which like most functionally related family members, produces mainly neoagarotetraose (LA-G-LA-G) and neoagarohexaose (LA-G-LA-G-LA-G) as final products from larger polymers [22] and is consistent with the open 30A long binding cleft of ZgAgaA [23], ZgAgaB, and ZgAgaD [22] (Figure 2a). Because smaller oligosaccharides with degrees of polymerization of four to six are refractory to hydrolysis by GH16 agarases, they represent intermediates during polymer hydrolysis and require
additional enzymes to be degraded in complete agarolytic systems. This function is performed by other b-agarases and a-agarases present in such systems, which are either GH50 b-agarases or GH117 a-agarases. Exo-processive GH50 b-agarase. The GH50 b-agarases produce either a neoagarobiose or neoagarotetraose product from their action on agarose. Aga50D from Saccharophagus degradans 2–40 releases neoagarobiose from the non-reducing end of neoagaro-oligosaccharides, thus continuing where GH16 b-agarases stop being active [24]. The structures of Aga50D, the first for the family, in complex with unhydrolyzed substrates and a disaccharide product revealed two domains, a catalytic domain with a (b/a)8-barrel fold and a domain resembling a b-jelly roll fold carbohydrate-binding module (CBM) (PDB 4BQ2) [25] (Figure 2b). Two catalytic glutamates, positioned in a way that suggests a retaining catalytic mechanism, are present at the bottom of a tunnel shaped active site, consistent with an exo-processive mode of action [26]. CBMs are typically connected with short flexible linkers to the catalytic domain, yet in Aga50D the CBM-like domain is fused with the catalytic domain at the entrance of the tunnel shaped active site, extending the substrate binding cleft by contributing a tryptophan residue to binding the substrate. a-Agarase, a neoagarobiose hydrolase in GH117. GH117 aagarases are exo-acting enzymes that remove 3,6-anhydro-L-galactose from the non-reducing end of neoagarooligosaccharides. The first structure of a GH117 a-agarase, from the marine bacterium Z. galactanivorans [27], was quickly followed by a structure of a GH117 enzyme from S. degradans 2–40 [28]. These structures revealed
Figure 2
(a)
(b)
(c)
(d)
Current Opinion in Structural Biology
Agars such as agarose and porphyran are degraded by enzymes from diverse GH families and folds. (a) Structure of endo-acting GH16 b-agarase ZgAgaD from marine bacterium Zobellia galactanivorans (PDB 4ASM). (b) Structure of the exo-acting GH50 b-agarase from Saccharuphagus degradans (Aga50D, PDB 4BQ5). (c) Monomer of the exo-acting GH117 a-neoagarobiose hydrolase from human gut bacterium Bacteroides plebeius (BpGH117, PDB 4AK7). (d) Structure of endo-acting b-porphyranase BpGH86B from B. plebeius. The structures are presented as cartoon plots, which are color ramped from the N-terminus (blue) to the C-terminus (red). The glycan ligands are shown as stick models. The figures were prepared with pymol, www.pymol.org. Current Opinion in Structural Biology 2014, 28:77–86
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Structures and mechanisms of marine CAZymes Hehemann, Boraston and Czjzek 81
these enzymes to have a five bladed b-propeller fold and to form stable dimers by domain swapping [27]. A third structure of a GH117 enzyme, BpGH117 (PDB 4AK5) from the human gut bacterium Bacteroides plebeius, was determined as a Michaelis-complex with neoagarobiose, confirming the presence of the active site at the center of the b-propeller and the proximity of the active site to a metal binding site, though the role of the metal binding site is unclear at this time [29] (Figure 2c). Interpretation of this structure in combination with site directed mutagenesis indicated an active site architecture most consistent with a catalytic mechanism that inverts the anomeric configuration. In the mechanism proposed for this enzyme a histidine side chain functions as the catalytic acid and an aspartate as the base. The histidine interacts closely with a separate aspartate, creating an Asp-His dyad similar to that of the retaining N-acetylglucosaminidase from Bacillus subtilis [30] and suggesting a relay system to deliver a proton to the leaving group. Curiously, many marine microbes contain multiple paralogous genes encoding several GH117, GH86, GH50 or GH16 enzymes, when only a minimal system comprising an endo-b-agarase, an exo-b-agarase, and an exo-a-agarase appears to be required for production of monosaccharides. This apparent redundancy raises the question of why bacteria frequently have multiple versions of these enzymes. Potentially, the frequent occurrence of multigenic families reflects an adaptation to the naturally occurring heterogeneity of agars, which may give rise to adaptive radiations in CAZyme subfamilies [22].
Porphyran degradation The first porphyranases were discovered in Z. galactanivorans and are members of family GH16 [8]. These enzymes are quite closely related to their agarolytic counterparts with the notable exception of their capacity to specifically recognize the sulfation pattern of porphyran (see following section). Remarkably, homologs of the genes in the porphyran degrading locus of Z. galactanivorans were found in the human gut bacterium B. plebeius [8], which became the source of the first GH86 b-porphyranase to be structurally and functionally characterized [31]. GH86 b-porphyranases. BpGH86A from B. plebeius has endo-b-porphyranase activity, the first such specificity for an enzyme in a family that was thought to be dominated by b-agarases. The enzyme adopts a (b/a)8-barrel fold with two ancillary b-sandwich domains (Figure 2d). The structure in complex with a product of porphyran hydrolysis, a hybrid-hexamer of agarose and porphyran (L6S-G-LA-G-L6S-G), revealed an arrangement of catalytic glutamates that was consistent with an anomeric configuration-retaining catalytic mechanism [26]. The 2 subsite of BpGH86 made specific interactions with a L6S residue and its 6-sulfate group, providing insight into the www.sciencedirect.com
selectivity of BpGH86A for porphyran. The gene encoding BpGH86 is found in a polysaccharide utilization locus together with a GH16 porphyranase and other agar active enzymes [31,29]. The biochemical, structural and functional studies of this gene cluster from B. plebeius have shown that the enzyme system is functional, actively enabling the bacterium to metabolize porphyran. The most closely related homologs of the enzymes found in the B. plebeius porphyran gene cluster are found in marine bacteria, supporting the hypothesis that functional carbohydrate degradation traits can be transferred from environmental bacteria to human gut microbes [8,31].
Sulfate-ester recognition in marine CAZymes — comparison across the different families A feature that is shared by many marine enzymes is their ability to specifically recognize the sulfate modifications that are abundant in the polysaccharides of marine algae. Figure 3 illustrates selected sulfate-ester binding pockets of various marine b-carrageenases (Figure 3a–d) and bporphyranases (Figure 3e–h). These structures exemplify that these enzymes have evolved through multiple changes in their active sites to adopt recognition of bulky and negatively charged sulfate-ester groups in a pocket of specific volume that is lined with a basic residue, usually an arginine. For example, in all enzyme-sulfated substrate complexes reported so far, except for the GH86 b-porphyranase of B. plebeius (Figure 3e,f), the guanidine-group of an arginine in the pocket hydrogen bonds with the sulfate group, providing specificity to the interaction. In BpGH86, the basic residues hydrogen bonding the sulfate groups are a histidine and a lysine in the different subsites. Another remarkable feature is that the catalytic grooves of these enzymes have, in general, fewer exposed hydrophobic surfaces than enzymes binding neutral polysaccharides but, paradoxically, the specific pockets accommodating the charged sulfate-group contain at least one hydrophobic residue (see Figure 3).
Lytic enzyme systems for digestion of brown seaweeds Brown seaweeds belong to the most productive algae in marine eco-systems. For example, kelp forests rapidly produce large amounts of biomass, forming a base of coastal food webs and constituting an important carbon sink [32]. Alginate consists of b-1,4-linked D-mannuronate (M) and its C5 epimer a-L-guluronate (G) and is the major cell wall polysaccharide of kelps (Figure 1). These monomers occur as homogeneous blocks of M or G, or as heteropolymeric blocks of alternating M and G units. Marine bacteria deconstruct alginate gels with alginate lyases and oligo-alginate lyases. These enzymes belong to the general class of PLs that utilize a b-elimination reaction to cleave the glycosidic bond. A description of selected marine polysaccharide-specific lyases is given below and a more comprehensive overview of structures and functions of PLs in general can be found in the Current Opinion in Structural Biology 2014, 28:77–86
(a)
(c) E245
H281
(g) W56
R260
R303
–2
–2 Y341 F271
+1
L273
(e)
G258
H53
W144
Y342
H53
–1
R133 S326
K394 Q424 (b)
Y341
W331
E163 (d)
D327 V134
(f)
(h) W78
W95
Q424
R70 W67
R151 I149
A342
–3
–2 Y83
W350
+3 R321
R187
–6 K87 W195
R252 I141 Current Opinion in Structural Biology
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Positively charged pockets that bind sulfate groups are common in enzymes that hydrolyze glycans from algae. (a,b) Clipped surface (colored in magenta) representation of substrate binding sites in the catalytic active site groove of GH82 iota-b-carrageenase (pdb code 1KTW) showing the specific pockets and residues accommodating the sulfate-ester functional groups in the +1 (panel a) and +3 (panel b) binding sites. (c,d) GH16 kappa-b-carrageenase (M Czjzek et al., unpublished) showing the specific pockets and residues accommodating the sulfate-ester functional groups in the 1 (panel c) and 3 (panel d) binding sites. (e,f) GH86 b-porphyranase (pdb code 4AW7) showing the specific pockets and residues accommodating the sulfate-ester functional groups in the 2 (panel e) and 6 (panel f) binding sites. (g) GH16 b-porphyranase (PorA) showing the specific pocket in the 2 binding site. (h) GH16 b-porphyranase (PorB) showing the binding site pocket accommodating a sulfate-ester containing buffer molecule (MES) in the 2 equivalent position. The carbohydrate sugar-units and the MES ligand in panel H are displayed as sticks, carbons are colored in white and oxygens in red. Specific residues surrounding the binding sites that form the pocket and interact with the ligand are also represented as sticks; carbons are colored in yellow, oxygens in red and nitrogen in blue. Hydrophobic residues within the sulfate binding pocket are also highlighted: these are L273 or A342 in the iota-carrageenase, F271 or I149 in the kappa-carrageenase, Y342 or Y83 in the b-porphyranase (GH86), V134 in GH16 PorA and I141 in PorB.
82 Carbohydrate–protein interactions and glycosylation
Current Opinion in Structural Biology 2014, 28:77–86
Figure 3
Structures and mechanisms of marine CAZymes Hehemann, Boraston and Czjzek 83
accompanying article by Garron and Cygler in this volume [33]. Alginate lyases. The structures of alginate lyases have been determined for examples from families PL7 [34], PL14 [35], PL15 [36], PL17 [37] (only post-2010 examples are cited; see the CAZy database for a complete census of all PL structures [5]). In general, PL7 enzymes play a similar role in alginate hydrolysis as that of GH16 enzymes in agar hydrolysis by initiating alginate degradation outside of the cell through endo-cleavage of alginate polysaccharide chains into shorter oligosaccharides, which are then substrates for the import systems [38–40]. PL7 lyases adopt a b-jellyroll fold with substrate binding clefts that span the entire length of the enzyme. However, some exceptions do exist as in the structure of an exo-acting PL7 that specifically degrades alginate has been described [34]. In this system, two enzymes belong to the PL7 family, AlyA1 (PDB 3ZPY) and AlyA5 (PDB 4BE3) from Z. galactanivorans, but they display complementary modes of action. AlyA1 is an endo-acting lyase with specificity for guluronate blocks and produces mainly trisaccharides and tetrasaccharides while AlyA5 has exo-activity and cleaves unsaturated monosaccharides from the non-reducing end of oligo-alginates and polysaccharide chains. Consistent with their mode of actions AlyA1 has an extended, open substrate binding cleft (Figure 4a), while the active site of AlyA5 forms a pocket; several characteristic loop insertions, present in the
sequence of AlyA5 wrap around the active site and close up one side of the cleft (Figure 4b) [34]. Following initial degradation of alginate by these PL7 enzymes, the smaller oligosaccharides are imported into the periplasmic space and further degraded by intracellular, exo-acting PL15 and PL17 oligoalginate lyases. PL15 & PL17 oligoalginate lyases. Oligoalginate lyases are crucial to complete alginate degradation as they process the oligosaccharides released by alginate lyases. The first structure of an oligoalginate lyase, Atu3025 from Agrobacterium tumefaciens, was reported in 2010 (PDB 3A0O) [36]. This PL15 enzyme adopts a bi-modular architecture with an (a/a)6-barrel domain and an anti-parallel b-sheet domain (Figure 4c). The structure of an inactive mutant of Atu3025 trapped with bound substrates (PDB 3AFL) revealed that it cleaves monomers from the non-reducing end of alginate chains in an active site pocket comprising three substrate binding subsites located at the interface between the two domains. The apo-enzyme (PDB 3A0O) has an open conformation but upon substrate binding undergoes a conformational change and folds over the substrate to form a pocket like active site. A conformational change of 108 locks the guluronate residue in the 1 subsite between two tyrosine residues. This movement allows a histidine residue and a tyrosine residue, which have been proposed to be the catalytic base and acid, respectively, to engage the sugar residue bound in the +1 subsite and the glycosidic bond, allowing the
Figure 4
(a)
(c)
(d)
(e)
(b)
Current Opinion in Structural Biology
Structural diversity of enzymes acting on the anionic cell wall polysaccharides alginate and ulvan from brown and green seaweeds. (a) Structure of endo-acting Pl7 alginate lyase AlyA1 (PDB 3ZPY) from marine bacterium Zobellia galactanivorans with open active site cleft. (b) Structure of Pl7 exoacing alginate lyase AlyA5 from Z. galactanivorans (PDB 4BE3) with a pocket shaped active site. (c) Structure of exo-acing Pl15 oligoalginate lyase from Agrobacterium tumefaciens (PDB 3AFL). (d) Structure of exo-acing Pl17 oligoalginate lyase from Saccharuphagus degradans (PDB 4OJZ). (e) Structure of GH105 a exo-acting b-glucuronyl hydrolase from Nonlabens ulvanivorans that degrades ulvan oligosaccharides (PDB 4CE7). The structures are presented as cartoon plots, which are color ramped from the N-terminus (blue), to the C-terminus (red). When present the glycan ligand are shown as stick models. The figures were prepared with pymol, www.pymol.org. www.sciencedirect.com
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84 Carbohydrate–protein interactions and glycosylation
correct positioning of the catalytic machinery and leading to lysis of the substrate [36]. The first structure of a PL17 enzyme, Alg17c from the marine bacterium S. degradans, has recently been solved (PDB 40JZ) [37]. Though sharing only about 10% sequence identity with terrestrial PL15 members, Alg17c adopts a similar (a/a)6-barrel + anti-parallel b-sheet (Figure 4d). However, based on the structure with substrate trapped in the active site of a Tyr to Ala mutant of Alg17c, different side chains, specifically two tyrosine residues, are proposed as catalytic the catalytic acid and base. Other notable differences in PL17 enzymes include the presence of a structural metal ion that is absent in PL15.
First enzymes for the degradation of green seaweed polysaccharides Ulva spp. are green seaweeds that grow attached to hard substrates or in a planktonic form for improved access to light and nutrients. The high growth rate and yields of these algae have an economic impact as they interfere with aquaculture or dissuade tourism, as when beaches are fouled with the abundant remnants of decomposing green seaweed. However, the high growth rate and yields that make these carbohydrate-rich algae a nuisance also suggest they may be valuable as feedstock for biofuels. The major matrix polysaccharide in Ulva spp. are called ulvans, which are anionic sulfated polysaccharides made up of 3-sulfated rhamnose (Rha3S), glucuronic acid (GlcA), iduronic acid (IduA), and some xylose (Xyl) units (Figure 1). The identification of the microbial enzyme systems responsible for their degradation will be key to unlocking the potential of using green algae as feedstocks. An ulvan degrading enzyme from GH105. A recent bioprospecting project in which bacteria capable of metabolizing ulvans were screened and isolated identified Nonlabens ulvanivorans from the feces of the marine gastropod Aplysia punctata as an efficient ulvan degrader. From the supernatant of this bacterium when grown in presence of ulvan two new ulvan lyases were identified, the genes encoding them cloned, and the recombinant gene products biochemically characterized [41]. Exploration of the genomic environment around the ulvan lyases revealed a gene highly similar to known unsaturated uronyl hydrolases classified in the CAZy glycoside hydrolase family 105. The crystal structure of this GH105 is the first of an ulvan degrading enzyme and showed it to have a (a/a)6 fold (Figure 4e). This unsaturated b-glucuronyl hydrolase acts on the oligosaccharides produced by the ulvan lyases [42]. Intriguingly, this enzyme displayed activity on bconfigured substrates, in contrast to all other enzymes so far characterized in this family. The authors argued that because recognition of the glycosidic bond is not part of the catalytic mechanism, lack of selection pressure for Current Opinion in Structural Biology 2014, 28:77–86
residues specific for either axial or equatorial configurations explains the presence of both specificities within one family. Indeed, in this case, the catalytic hydrolysis occurs through water addition to the C4–C5 unsaturated bond, followed by molecular rearrangements that ultimately lead to the cleavage of the glycosidic bond. Once again, this example demonstrates that with the increasing numbers of GH families having structural representatives and well characterized members, the number of ‘exceptions’ to established patterns or rules is also increasing [43,44].
Conclusion The growing ecological and biotechnological interest in algal biomass conversion has led to an increased effort towards the biochemical and structural characterization of marine CAZymes. For some major marine polysaccharides, such as agarose or alginate, these recent advances have provided insight into all enzymatic steps leading from polysaccharide to monosaccharides. However, recent genomic and meta-genomic data suggest that we are far from having covered the diversity and complexity of marine carbohydrate-degrading systems. In particular, oligosaccharide and polysaccharide degrading enzymes are still to be identified for many algal polysaccharides, especially the ones produced by marine microalgae. A metagenomic and proteomic survey of the dynamics of a microalgae bloom and associated bacterial communities identified >20 bacterial GH and PL families and associated enzymes, such as sulfatases, all of which were most likely used by the different bacterial phyla to digest and consume the algal polysaccharides that became successively available during the algal bloom [12]. However, the substrates of most of the putative CAZymes were unknown highlighting the need to characterize these enzymes to understand their precise role within the marine carbon cycle.
Acknowledgements ABB is thankful for the support of a Natural Sciences and Engineering Research Council of Canada Discovery Grant and an E.W.R. Steacie Memorial Fellowship. JHH was funded by an EMBO Long Term Fellowship and is thankful to receive a fellowship from the Human Frontiers Program. MC is thankful for support by the French Centre National de Recherches Scientifiques. This work also benefited from the support of the French Government through the National Research Agency with regards to an investment expenditure program IDEALG which reference is stated as ANR-10-BTBR-04.
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29. Hehemann J-H, Smyth L, Yadav A, Vocadlo DJ, Boraston AB: Analysis of keystone enzyme in agar hydrolysis provides insight into the degradation (of a polysaccharide from) red seaweeds. J Biol Chem 2012, 287:13985-13995. Crystal structure of a GH117 a-agarase with a Michaelis-complex of the substrate neoagarobiose reveals a histidine as the catalytic acid and which forms a catalytic dyad with an adjacent aspartate residue suggesting a proton relay system to the leaving group. 30. Litzinger S, Fischer S, Polzer P, Diederichs K, Welte W, Mayer C: Structural and kinetic analysis of Bacillus subtilis Nacetylglucosaminidase reveals a unique Asp-His dyad mechanism. J Biol Chem 2010, 285:35675-35684. 31. Hehemann J-H, Kelly AG, Pudlo NA, Martens EC, Boraston AB: Bacteria of the human gut microbiome catabolize red seaweed glycans with carbohydrate-active enzyme updates from extrinsic microbes. Proc Natl Acad Sci U S A 2012, 109:19786-19791. Describes a high-resolution family-first crystal structure of a GH86 porphyranase that originates from a human gut bacterium. The enzyme is involved in the degradation of the sulfated polysaccharide porphyran from the red seaweed Nori. Current Opinion in Structural Biology 2014, 28:77–86
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ScienceDirect A sweet new wave: structures and mechanisms of enzymes that digest polysaccharides from marine algae Jan-Hendrik Hehemann1, Alisdair B Boraston2 and Mirjam Czjzek3,4 Marine algae contribute approximately half of the global primary production. The large amounts of polysaccharides synthesized by these algae are degraded and consumed by microbes that utilize carbohydrate-active enzymes (CAZymes), thus creating one of the largest and most dynamic components of the Earth’s carbon cycle. Over the last decade, structural and functional characterizations of marine CAZymes have revealed a diverse set of scaffolds and mechanisms that are used to degrade agars, carrageenan, alginate and ulvanpolysaccharides from red, brown and green seaweeds, respectively. The analysis of these CAZymes is not only expanding our understanding of their functions but is enabling the enhanced annotation of (meta)-genomic data sets, thus promoting an improved understanding of microbes that drive this marine component of the carbon cycle. Furthermore, this information is setting a foundation that will enable marine algae to be harnessed as a novel resource for biorefineries. In this review, we cover the most recent structural and functional analyses of marine CAZymes that are specialized in the digestion of macro-algal polysaccharides. Addresses 1 Massachusetts Institute of Technology, Department of Civil and Environmental Engineering, 15 Vassar Street, Bldg 48-108, Cambridge, MA 02139, USA 2 Biochemistry and Microbiology, University of Victoria, PO Box 3055 STN CSC, Victoria, BC, Canada V8W 3P6 3 Sorbonne Universite´s, UPMC, Universite´ Paris 06, France 4 CNRS UMR 8227, Integrative Biology of Marine Models, Station Biologique de Roscoff, CS 90074, F-29688 Roscoff cedex, Bretagne, France Corresponding authors: Hehemann, Jan-Hendrik ([email protected]), Czjzek, Mirjam ([email protected])
Current Opinion in Structural Biology 2014, 28:77–86 This review comes from a themed issue on Carbohydrate–protein interactions and glycosylation Edited by Harry Brumer and Harry J Gilbert
http://dx.doi.org/10.1016/j.sbi.2014.07.009 0959-440X/# 2014 Elsevier Ltd. All rights reserved.
Introduction About one-half of the global net primary production, which amounts to an estimated 104.9 pentagrams of www.sciencedirect.com
carbon per year, is generated by marine algae [1]. Marine heterotrophic microbes digest the produced organic matter, such as high molecular weight algal polysaccharides, into low molecular weight substrates. This is amongst the largest and fastest catabolic biotransformations on earth and is key to the carbon cycle by returning this photosynthetically fixed carbon back to the atmosphere. Indeed, the ocean has been compared to ‘a global digester, where almost all organic matter injected from surface waters (0–100 m) is respired to inorganic nutrients’ [2] and only a ‘small’ fraction escapes rapid microbial consumption. That this phenomenon exists and is critical to both ocean health and earth climate regulation is well appreciated; however, the microbial processes governing this turnover are not well understood, particularly at the molecular level. For example, on average, marine genomic or metagenomic data sets contain genes encoding up to 20% more hypothetical proteins with unknown functions and structures, or ‘orphans’, than their terrestrial counterparts [3]. Countless marine microorganisms associated with algae contain unexplored enzymes, such as novel glycoside hydrolases (GH) and polysaccharide lyases (PL), that participate in important metabolic pathways and whose characterization holds the potential to provide new insights into marine carbon recycling. The CAZy database (www.cazy.org) groups carbohydrateactive enzymes (CAZymes) that have a common ancestor into families of proteins and presently includes 133 GH families and 33 PL families [4,5]. Recent years have seen increasing interest in the analysis of CAZymes that degrade marine polysaccharides, mainly those from seaweeds. The first structure determined for such an enzyme was the kappa-carrageenase from the marine bacterium Pseudoalteromonas carrageenovora, which was isolated for its capacity to grow on carrageenan-gels [6]. While this enzyme belongs to the large, multi-specific family GH16, in which both ‘terrestrial’ and ‘marine’ enzymes occur [7,6,8,9,10], many more recently identified marine polysaccharide specific CAZymes have laid the basis for several novel and marine-dominant GH and PL families, such as b-agarases in GH50, GH86, and GH118; i-carrageenase in GH82; aagarases in GH96 and GH117; and alginate lyases in PL7, PL15, and PL17. As the database of CAZymes becomes increasingly populated with known and putative enzymes involved in the breakdown of marine algal polysaccharides some of the important questions directing the examination of these Current Opinion in Structural Biology 2014, 28:77–86
78 Carbohydrate–protein interactions and glycosylation
Table 1 CAZy famillies containing enzymes active on marine polysaccharides and their respective PDB accession codes CAZy family GH16
Protein
GH50
k-Carrageenase (Pseudoaltermonas carragenovora) b-Agarase A (Zobellia galactanivorans) b-Agarase B (Zobellia galactanivorans) b-Agarase D (Zobellia galactanivorans) b-Porphyranase (Bacteroides plebeius) b-Porphyranase A (Zobellia galactanivorans) b-Porphyranase B (Zobellia galactanivorans) b-1,3-Glucanase (Mycobacterium marinum M) Laminarinase (Thermotoga maritima) Laminarinase R (Rhodothermus marinus) Laminarinase A (Pyrococcus furiosus) Laminarinase A (Zobellia galactanivorans) Exo-b-agarase D (Saccharophagus degradans)
GH82
i-Carrageenase (Alteromonas fortis)
GH86 GH105 GH117
b-Porphyranase (Bacteroides plebeius) Unsaturated b-glucuronyl hydrolase (Nonlabens ulvanivorans) a-1,3-L-(3,6-Anhydro)-galactosidase (Zobellia galactanivorans) a-1,3-L-Neoagarobiose hydrolase (Saccharophagus degradans) a-1,3-L-(3,6-Anhydro)-galactosidase (Bacteroides plebeius) Endo-guluronate lyase (Zobellia galactanivorans) Exo-alginate lyase (Zobellia galactanivorans) Exo-alginate lyase (Agrobacterium fabrum) Oligoalginate lyase (Saccharophagus degradans)
PL7 PL15 PL17
PDB codes 1DYP 1O4Y; 1URX a 1O4Z; 4ATF 4ASM 4AWD 4ATE; 3ILF 3JUU 4PQ9 3AZX; 3AZZ 3ILN 2VY0 4BQ1; 4BOW 4BQ2; 4BQ3 4BQ4; 4BQ5 1H80; 1KTW 3LMW 4AW7 4CE7 3P2N 3R4Y; 3R4Z 4AK5; 4AK7 3ZPY 4BE3 3A0O; 3AFL 4NEI; 4OJZ
A complete overview of the structural state of all CAZyme families so far can be found in Lombard et al. [5]. For all GH16 enzymes, only one additional pdb code of a substrate-complex is given, when existent.
a
enzymes are: What are the precise specificities of these enzymes? What is the molecular basis behind enzymatic recognition of the unique structures of marine polysaccharides? What is the biochemistry underlying the enzymatic cleavage of glycosidic bonds in marine polysaccharides? The answers to these questions are helping us to understand how microbes process the billions of tons of polysaccharides that are produced annually by algae in the sea and informing how this process may change in future marine systems [11]. Importantly, such research is also providing biotechnological tools to produce tailored oligosaccharides with therapeutic potential and generating the biocatalysts required for efficient use of algal biomass as feedstocks for biorefinement processes [12]. Here we provide an update on the most recent advances in understanding enzymes involved in marine polysaccharide degradation (Table 1).
Seaweed degrading CAZymes for biorefinery applications In the quest for alternative biofuels seaweeds are a promising compliment to, or even substitute for, land plant biomass as a feedstock. Algae grow faster and do not impinge on arable land. Furthermore, seaweeds build their cell walls with higher amounts of gel forming polysaccharides that are highly hydrated and, relative to common ligno-cellulosic biomass, much easier to digest with enzymes. For example, depending on the type of seaweed, the cell walls of these macro-algae (Figure 1) Current Opinion in Structural Biology 2014, 28:77–86
contain up to 40% agarose, porphyran, carrageenan, alginate, or ulvan polysaccharides, most of which form hydrogels [13,14]. Recently, this advantage was successfully exploited by transferring an alginate-specific metabolic locus from Vibrio splendidus 12B01 [15] into a bioengineered Escherichia coli strain, resulting in a bacterium able to convert alginates from brown seaweeds into high yields of bioethanol [16]. This example illustrates the great potential of marine polysaccharide specific CAZymes for bioenergy production. Optimization of this approach to convert algal alginate into biofuel, however, and extension of the concept to utilize seaweeds rich in other polysaccharides requires ongoing structural and functional characterization of algae degrading GHs and PLs from marine microbes.
Hydrolytic enzyme systems for the digestion of red seaweeds The matrix polysaccharides of red algae are either agars (in agarophytes) or carrageenan (in carragenophytes). Enzymes degrading the latter polysaccharide were discovered early on during CAZyme prospecting and are extensively reviewed elsewhere [17]. Although bagarases have been characterized for some time, their structural analysis is a relatively recent development, as is the discovery and characterization of a-agarases. Agars are galactans comprising L-galactose that is a-1,3 linked with D-galactose (G). In agarose, the L-galactose is www.sciencedirect.com
Structures and mechanisms of marine CAZymes Hehemann, Boraston and Czjzek 79
Figure 1
2 1
3
4
3. GH117
1. GH16 OH HO
O
2. GH50
O
O
agarose
O
OH
O
O OH OHOH
GH16, GH86
OH
OH O HO
O O
porphyran
OH OH
O
O
O
OH
HO OH
O
O
OH
O O
OH
O
O
O
O
–O3S
OH
O –O3S
O
OH
1. PL7 OH
–OOC O
O OH
–OOC O HO
O OH
O
2. PL15, PL17, PL7 AlyA5 OH O
HO O
O OH
alginate
O COO-
–OOC OH
–OOC
GH105
–OOC O HO OH
H 3C O
OH
O
ulvan O
–O3S
O O
O HO
H3C O
O O
–O3S
OH
OH Current Opinion in Structural Biology
Red, green and brown algae contain different types of polysaccharides that are degraded by dedicated enzymatic systems. Left: Coast of rocky beach near Roscoff (Brittany, France) at low tide shows different types of seaweeds. Most visible are brown seaweeds of Fucus spp. and sugar kelp Saccharina lattissima Right: Close-up of left showing a patch with diversity of seaweeds present in this ecosystem. Inset legend: (1,2) the red seaweeds Porphyra spp. and Chondrus spp. respectively; (3) the green seaweed Ulva spp.; (4) the brown seaweed Fucus spp. The major cell wall matrix polysaccharides are shown. Agarose and porphyran are abundant in red seaweeds, alginate is present in brown seaweeds and ulvan comes from green seaweeds. These polysaccharides are degraded by glycoside hydrolases and polysaccharide lyases belonging to different families as indicated above the glycosidic bonds. The sequence in which these enzymes operate is indicated by numbers. For example, in alginate degradation the PL15, PL17 and PL7AlyA5 remove unsaturated monosaccharides from the non-reducing end of oligosaccharides produced by endo-acting Pl7 alginate lyases that initiate digestion.
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80 Carbohydrate–protein interactions and glycosylation
3,6-anhydro-L-galactose (LA) while in porphyran it is Lgalactose-6-sulfate (L6S). The resulting neoagarobiose and neoporphyrobiose disaccharides are connected by b-1,4 glycosidic linkages in agarose and porphyran, respectively (Figure 1) [18]. To degrade these polysaccharides marine microbes use a variety of enzymes to specifically hydrolyze the b-glycosidic and a-glycosidic linkages. Those that have been structurally characterized are covered in more detail below and discussed within the context of agarolytic systems.
Agarose degradation At present, b-agarases belong to families GH16, GH50, GH86, and GH118, while and a-agarases belong to families GH96 and GH117. Those that are structurally characterized are in GH16, GH50, and GH117. b-Agarases in GH16. The first three-dimensional structures of two b-agarases were of the enzymes ZgAgaA and ZgAgaB, which originate from the marine bacterium Zobellia galactanivorans. These enzymes belong to family GH16, have a b-jelly-roll-fold, and catalyze hydrolysis with a catalytic mechanism that retains the stereochemistry at the anomeric carbon and utilizes two catalytic glutamate residues [19–21]. Their endo-mode of action, which like most functionally related family members, produces mainly neoagarotetraose (LA-G-LA-G) and neoagarohexaose (LA-G-LA-G-LA-G) as final products from larger polymers [22] and is consistent with the open 30A long binding cleft of ZgAgaA [23], ZgAgaB, and ZgAgaD [22] (Figure 2a). Because smaller oligosaccharides with degrees of polymerization of four to six are refractory to hydrolysis by GH16 agarases, they represent intermediates during polymer hydrolysis and require
additional enzymes to be degraded in complete agarolytic systems. This function is performed by other b-agarases and a-agarases present in such systems, which are either GH50 b-agarases or GH117 a-agarases. Exo-processive GH50 b-agarase. The GH50 b-agarases produce either a neoagarobiose or neoagarotetraose product from their action on agarose. Aga50D from Saccharophagus degradans 2–40 releases neoagarobiose from the non-reducing end of neoagaro-oligosaccharides, thus continuing where GH16 b-agarases stop being active [24]. The structures of Aga50D, the first for the family, in complex with unhydrolyzed substrates and a disaccharide product revealed two domains, a catalytic domain with a (b/a)8-barrel fold and a domain resembling a b-jelly roll fold carbohydrate-binding module (CBM) (PDB 4BQ2) [25] (Figure 2b). Two catalytic glutamates, positioned in a way that suggests a retaining catalytic mechanism, are present at the bottom of a tunnel shaped active site, consistent with an exo-processive mode of action [26]. CBMs are typically connected with short flexible linkers to the catalytic domain, yet in Aga50D the CBM-like domain is fused with the catalytic domain at the entrance of the tunnel shaped active site, extending the substrate binding cleft by contributing a tryptophan residue to binding the substrate. a-Agarase, a neoagarobiose hydrolase in GH117. GH117 aagarases are exo-acting enzymes that remove 3,6-anhydro-L-galactose from the non-reducing end of neoagarooligosaccharides. The first structure of a GH117 a-agarase, from the marine bacterium Z. galactanivorans [27], was quickly followed by a structure of a GH117 enzyme from S. degradans 2–40 [28]. These structures revealed
Figure 2
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Agars such as agarose and porphyran are degraded by enzymes from diverse GH families and folds. (a) Structure of endo-acting GH16 b-agarase ZgAgaD from marine bacterium Zobellia galactanivorans (PDB 4ASM). (b) Structure of the exo-acting GH50 b-agarase from Saccharuphagus degradans (Aga50D, PDB 4BQ5). (c) Monomer of the exo-acting GH117 a-neoagarobiose hydrolase from human gut bacterium Bacteroides plebeius (BpGH117, PDB 4AK7). (d) Structure of endo-acting b-porphyranase BpGH86B from B. plebeius. The structures are presented as cartoon plots, which are color ramped from the N-terminus (blue) to the C-terminus (red). The glycan ligands are shown as stick models. The figures were prepared with pymol, www.pymol.org. Current Opinion in Structural Biology 2014, 28:77–86
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Structures and mechanisms of marine CAZymes Hehemann, Boraston and Czjzek 81
these enzymes to have a five bladed b-propeller fold and to form stable dimers by domain swapping [27]. A third structure of a GH117 enzyme, BpGH117 (PDB 4AK5) from the human gut bacterium Bacteroides plebeius, was determined as a Michaelis-complex with neoagarobiose, confirming the presence of the active site at the center of the b-propeller and the proximity of the active site to a metal binding site, though the role of the metal binding site is unclear at this time [29] (Figure 2c). Interpretation of this structure in combination with site directed mutagenesis indicated an active site architecture most consistent with a catalytic mechanism that inverts the anomeric configuration. In the mechanism proposed for this enzyme a histidine side chain functions as the catalytic acid and an aspartate as the base. The histidine interacts closely with a separate aspartate, creating an Asp-His dyad similar to that of the retaining N-acetylglucosaminidase from Bacillus subtilis [30] and suggesting a relay system to deliver a proton to the leaving group. Curiously, many marine microbes contain multiple paralogous genes encoding several GH117, GH86, GH50 or GH16 enzymes, when only a minimal system comprising an endo-b-agarase, an exo-b-agarase, and an exo-a-agarase appears to be required for production of monosaccharides. This apparent redundancy raises the question of why bacteria frequently have multiple versions of these enzymes. Potentially, the frequent occurrence of multigenic families reflects an adaptation to the naturally occurring heterogeneity of agars, which may give rise to adaptive radiations in CAZyme subfamilies [22].
Porphyran degradation The first porphyranases were discovered in Z. galactanivorans and are members of family GH16 [8]. These enzymes are quite closely related to their agarolytic counterparts with the notable exception of their capacity to specifically recognize the sulfation pattern of porphyran (see following section). Remarkably, homologs of the genes in the porphyran degrading locus of Z. galactanivorans were found in the human gut bacterium B. plebeius [8], which became the source of the first GH86 b-porphyranase to be structurally and functionally characterized [31]. GH86 b-porphyranases. BpGH86A from B. plebeius has endo-b-porphyranase activity, the first such specificity for an enzyme in a family that was thought to be dominated by b-agarases. The enzyme adopts a (b/a)8-barrel fold with two ancillary b-sandwich domains (Figure 2d). The structure in complex with a product of porphyran hydrolysis, a hybrid-hexamer of agarose and porphyran (L6S-G-LA-G-L6S-G), revealed an arrangement of catalytic glutamates that was consistent with an anomeric configuration-retaining catalytic mechanism [26]. The 2 subsite of BpGH86 made specific interactions with a L6S residue and its 6-sulfate group, providing insight into the www.sciencedirect.com
selectivity of BpGH86A for porphyran. The gene encoding BpGH86 is found in a polysaccharide utilization locus together with a GH16 porphyranase and other agar active enzymes [31,29]. The biochemical, structural and functional studies of this gene cluster from B. plebeius have shown that the enzyme system is functional, actively enabling the bacterium to metabolize porphyran. The most closely related homologs of the enzymes found in the B. plebeius porphyran gene cluster are found in marine bacteria, supporting the hypothesis that functional carbohydrate degradation traits can be transferred from environmental bacteria to human gut microbes [8,31].
Sulfate-ester recognition in marine CAZymes — comparison across the different families A feature that is shared by many marine enzymes is their ability to specifically recognize the sulfate modifications that are abundant in the polysaccharides of marine algae. Figure 3 illustrates selected sulfate-ester binding pockets of various marine b-carrageenases (Figure 3a–d) and bporphyranases (Figure 3e–h). These structures exemplify that these enzymes have evolved through multiple changes in their active sites to adopt recognition of bulky and negatively charged sulfate-ester groups in a pocket of specific volume that is lined with a basic residue, usually an arginine. For example, in all enzyme-sulfated substrate complexes reported so far, except for the GH86 b-porphyranase of B. plebeius (Figure 3e,f), the guanidine-group of an arginine in the pocket hydrogen bonds with the sulfate group, providing specificity to the interaction. In BpGH86, the basic residues hydrogen bonding the sulfate groups are a histidine and a lysine in the different subsites. Another remarkable feature is that the catalytic grooves of these enzymes have, in general, fewer exposed hydrophobic surfaces than enzymes binding neutral polysaccharides but, paradoxically, the specific pockets accommodating the charged sulfate-group contain at least one hydrophobic residue (see Figure 3).
Lytic enzyme systems for digestion of brown seaweeds Brown seaweeds belong to the most productive algae in marine eco-systems. For example, kelp forests rapidly produce large amounts of biomass, forming a base of coastal food webs and constituting an important carbon sink [32]. Alginate consists of b-1,4-linked D-mannuronate (M) and its C5 epimer a-L-guluronate (G) and is the major cell wall polysaccharide of kelps (Figure 1). These monomers occur as homogeneous blocks of M or G, or as heteropolymeric blocks of alternating M and G units. Marine bacteria deconstruct alginate gels with alginate lyases and oligo-alginate lyases. These enzymes belong to the general class of PLs that utilize a b-elimination reaction to cleave the glycosidic bond. A description of selected marine polysaccharide-specific lyases is given below and a more comprehensive overview of structures and functions of PLs in general can be found in the Current Opinion in Structural Biology 2014, 28:77–86
(a)
(c) E245
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D327 V134
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R252 I141 Current Opinion in Structural Biology
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Positively charged pockets that bind sulfate groups are common in enzymes that hydrolyze glycans from algae. (a,b) Clipped surface (colored in magenta) representation of substrate binding sites in the catalytic active site groove of GH82 iota-b-carrageenase (pdb code 1KTW) showing the specific pockets and residues accommodating the sulfate-ester functional groups in the +1 (panel a) and +3 (panel b) binding sites. (c,d) GH16 kappa-b-carrageenase (M Czjzek et al., unpublished) showing the specific pockets and residues accommodating the sulfate-ester functional groups in the 1 (panel c) and 3 (panel d) binding sites. (e,f) GH86 b-porphyranase (pdb code 4AW7) showing the specific pockets and residues accommodating the sulfate-ester functional groups in the 2 (panel e) and 6 (panel f) binding sites. (g) GH16 b-porphyranase (PorA) showing the specific pocket in the 2 binding site. (h) GH16 b-porphyranase (PorB) showing the binding site pocket accommodating a sulfate-ester containing buffer molecule (MES) in the 2 equivalent position. The carbohydrate sugar-units and the MES ligand in panel H are displayed as sticks, carbons are colored in white and oxygens in red. Specific residues surrounding the binding sites that form the pocket and interact with the ligand are also represented as sticks; carbons are colored in yellow, oxygens in red and nitrogen in blue. Hydrophobic residues within the sulfate binding pocket are also highlighted: these are L273 or A342 in the iota-carrageenase, F271 or I149 in the kappa-carrageenase, Y342 or Y83 in the b-porphyranase (GH86), V134 in GH16 PorA and I141 in PorB.
82 Carbohydrate–protein interactions and glycosylation
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Figure 3
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accompanying article by Garron and Cygler in this volume [33]. Alginate lyases. The structures of alginate lyases have been determined for examples from families PL7 [34], PL14 [35], PL15 [36], PL17 [37] (only post-2010 examples are cited; see the CAZy database for a complete census of all PL structures [5]). In general, PL7 enzymes play a similar role in alginate hydrolysis as that of GH16 enzymes in agar hydrolysis by initiating alginate degradation outside of the cell through endo-cleavage of alginate polysaccharide chains into shorter oligosaccharides, which are then substrates for the import systems [38–40]. PL7 lyases adopt a b-jellyroll fold with substrate binding clefts that span the entire length of the enzyme. However, some exceptions do exist as in the structure of an exo-acting PL7 that specifically degrades alginate has been described [34]. In this system, two enzymes belong to the PL7 family, AlyA1 (PDB 3ZPY) and AlyA5 (PDB 4BE3) from Z. galactanivorans, but they display complementary modes of action. AlyA1 is an endo-acting lyase with specificity for guluronate blocks and produces mainly trisaccharides and tetrasaccharides while AlyA5 has exo-activity and cleaves unsaturated monosaccharides from the non-reducing end of oligo-alginates and polysaccharide chains. Consistent with their mode of actions AlyA1 has an extended, open substrate binding cleft (Figure 4a), while the active site of AlyA5 forms a pocket; several characteristic loop insertions, present in the
sequence of AlyA5 wrap around the active site and close up one side of the cleft (Figure 4b) [34]. Following initial degradation of alginate by these PL7 enzymes, the smaller oligosaccharides are imported into the periplasmic space and further degraded by intracellular, exo-acting PL15 and PL17 oligoalginate lyases. PL15 & PL17 oligoalginate lyases. Oligoalginate lyases are crucial to complete alginate degradation as they process the oligosaccharides released by alginate lyases. The first structure of an oligoalginate lyase, Atu3025 from Agrobacterium tumefaciens, was reported in 2010 (PDB 3A0O) [36]. This PL15 enzyme adopts a bi-modular architecture with an (a/a)6-barrel domain and an anti-parallel b-sheet domain (Figure 4c). The structure of an inactive mutant of Atu3025 trapped with bound substrates (PDB 3AFL) revealed that it cleaves monomers from the non-reducing end of alginate chains in an active site pocket comprising three substrate binding subsites located at the interface between the two domains. The apo-enzyme (PDB 3A0O) has an open conformation but upon substrate binding undergoes a conformational change and folds over the substrate to form a pocket like active site. A conformational change of 108 locks the guluronate residue in the 1 subsite between two tyrosine residues. This movement allows a histidine residue and a tyrosine residue, which have been proposed to be the catalytic base and acid, respectively, to engage the sugar residue bound in the +1 subsite and the glycosidic bond, allowing the
Figure 4
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Structural diversity of enzymes acting on the anionic cell wall polysaccharides alginate and ulvan from brown and green seaweeds. (a) Structure of endo-acting Pl7 alginate lyase AlyA1 (PDB 3ZPY) from marine bacterium Zobellia galactanivorans with open active site cleft. (b) Structure of Pl7 exoacing alginate lyase AlyA5 from Z. galactanivorans (PDB 4BE3) with a pocket shaped active site. (c) Structure of exo-acing Pl15 oligoalginate lyase from Agrobacterium tumefaciens (PDB 3AFL). (d) Structure of exo-acing Pl17 oligoalginate lyase from Saccharuphagus degradans (PDB 4OJZ). (e) Structure of GH105 a exo-acting b-glucuronyl hydrolase from Nonlabens ulvanivorans that degrades ulvan oligosaccharides (PDB 4CE7). The structures are presented as cartoon plots, which are color ramped from the N-terminus (blue), to the C-terminus (red). When present the glycan ligand are shown as stick models. The figures were prepared with pymol, www.pymol.org. www.sciencedirect.com
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84 Carbohydrate–protein interactions and glycosylation
correct positioning of the catalytic machinery and leading to lysis of the substrate [36]. The first structure of a PL17 enzyme, Alg17c from the marine bacterium S. degradans, has recently been solved (PDB 40JZ) [37]. Though sharing only about 10% sequence identity with terrestrial PL15 members, Alg17c adopts a similar (a/a)6-barrel + anti-parallel b-sheet (Figure 4d). However, based on the structure with substrate trapped in the active site of a Tyr to Ala mutant of Alg17c, different side chains, specifically two tyrosine residues, are proposed as catalytic the catalytic acid and base. Other notable differences in PL17 enzymes include the presence of a structural metal ion that is absent in PL15.
First enzymes for the degradation of green seaweed polysaccharides Ulva spp. are green seaweeds that grow attached to hard substrates or in a planktonic form for improved access to light and nutrients. The high growth rate and yields of these algae have an economic impact as they interfere with aquaculture or dissuade tourism, as when beaches are fouled with the abundant remnants of decomposing green seaweed. However, the high growth rate and yields that make these carbohydrate-rich algae a nuisance also suggest they may be valuable as feedstock for biofuels. The major matrix polysaccharide in Ulva spp. are called ulvans, which are anionic sulfated polysaccharides made up of 3-sulfated rhamnose (Rha3S), glucuronic acid (GlcA), iduronic acid (IduA), and some xylose (Xyl) units (Figure 1). The identification of the microbial enzyme systems responsible for their degradation will be key to unlocking the potential of using green algae as feedstocks. An ulvan degrading enzyme from GH105. A recent bioprospecting project in which bacteria capable of metabolizing ulvans were screened and isolated identified Nonlabens ulvanivorans from the feces of the marine gastropod Aplysia punctata as an efficient ulvan degrader. From the supernatant of this bacterium when grown in presence of ulvan two new ulvan lyases were identified, the genes encoding them cloned, and the recombinant gene products biochemically characterized [41]. Exploration of the genomic environment around the ulvan lyases revealed a gene highly similar to known unsaturated uronyl hydrolases classified in the CAZy glycoside hydrolase family 105. The crystal structure of this GH105 is the first of an ulvan degrading enzyme and showed it to have a (a/a)6 fold (Figure 4e). This unsaturated b-glucuronyl hydrolase acts on the oligosaccharides produced by the ulvan lyases [42]. Intriguingly, this enzyme displayed activity on bconfigured substrates, in contrast to all other enzymes so far characterized in this family. The authors argued that because recognition of the glycosidic bond is not part of the catalytic mechanism, lack of selection pressure for Current Opinion in Structural Biology 2014, 28:77–86
residues specific for either axial or equatorial configurations explains the presence of both specificities within one family. Indeed, in this case, the catalytic hydrolysis occurs through water addition to the C4–C5 unsaturated bond, followed by molecular rearrangements that ultimately lead to the cleavage of the glycosidic bond. Once again, this example demonstrates that with the increasing numbers of GH families having structural representatives and well characterized members, the number of ‘exceptions’ to established patterns or rules is also increasing [43,44].
Conclusion The growing ecological and biotechnological interest in algal biomass conversion has led to an increased effort towards the biochemical and structural characterization of marine CAZymes. For some major marine polysaccharides, such as agarose or alginate, these recent advances have provided insight into all enzymatic steps leading from polysaccharide to monosaccharides. However, recent genomic and meta-genomic data suggest that we are far from having covered the diversity and complexity of marine carbohydrate-degrading systems. In particular, oligosaccharide and polysaccharide degrading enzymes are still to be identified for many algal polysaccharides, especially the ones produced by marine microalgae. A metagenomic and proteomic survey of the dynamics of a microalgae bloom and associated bacterial communities identified >20 bacterial GH and PL families and associated enzymes, such as sulfatases, all of which were most likely used by the different bacterial phyla to digest and consume the algal polysaccharides that became successively available during the algal bloom [12]. However, the substrates of most of the putative CAZymes were unknown highlighting the need to characterize these enzymes to understand their precise role within the marine carbon cycle.
Acknowledgements ABB is thankful for the support of a Natural Sciences and Engineering Research Council of Canada Discovery Grant and an E.W.R. Steacie Memorial Fellowship. JHH was funded by an EMBO Long Term Fellowship and is thankful to receive a fellowship from the Human Frontiers Program. MC is thankful for support by the French Centre National de Recherches Scientifiques. This work also benefited from the support of the French Government through the National Research Agency with regards to an investment expenditure program IDEALG which reference is stated as ANR-10-BTBR-04.
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