Functional characterization of a novel “ulvan utilization loci” found in Alteromonas sp. LOR genome

Algal Research 25 (2017) 39–46

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Functional characterization of a novel “ulvan utilization loci” found in Alteromonas sp. LOR genome

MARK

Elizabeth Forana,1, Vitaliy Buravenkova,1, Moran Kopela, Naama Mizrahia, Sivan Shoshania, William Helbertb,⁎, Ehud Banina,⁎ a

The Institute for Nanotechnology and Advanced Materials, The Mina and Everard Goodman Faculty of Life Sciences, Bar-Ilan University, Ramat Gan 52900, Israel Centre de Recherches sur les Macromolécules Végétales (UPR-CNRS 5301), Université Joseph Fourier and Institut de Chimie Moléculaire de Grenoble (ICMG, FR-CNRS 2607), Grenoble Cedex 9, France

b

A R T I C L E I N F O

A B S T R A C T

Keywords: Ulvan Polysaccharide Lyases Bacteria

Green algae belonging to the genus Ulvales are known to produce ulvan which is one of the main polysaccharide components of their cell wall. Ulvan is composed of 3O-sulfate-rhamnose (Rha3S), glucuronic acid (GlcA), iduronic acid (IduA) and xylose (Xyl) distributed in three disaccharide repetition moieties: [→4)-β-D-GlcA-(1 → 4)-α-L-Rha3S-(1 →], [→ 4)-α-L-IdoA-(1 → 4)-α-L-Rha3S(1 →] and [→ 4)-β-D-Xyl-(1 → 4)-α-L-Rha3S(1 →]. The ability of bacteria to degrade complex algal polysaccharides such as ulvan is usually encoded in clusters of genes referred to as polysaccharide utilization loci (PUL). Full saccharification of ulvan is expected to require an ulvan lyase, which cleaves the β-(1 → 4)-glycosidic bond between Rha3S and GluA or IduA through a β-elimination mechanism. In addition, enzymes with β-glucuronyl hydrolase, rhamnosidase, xylosidase and sulfatase activity are also expected. Recently, the genomes of several ulvan degrading bacteria were sequenced, which led to the identification of a new family of polysaccharide lyases family 24 (PL24). In this work, we have continued to mine the genomic data of one of the sequenced strains, Alteromonas sp. LOR. Here we report the identification of an ulvan associated PUL residing between open reading frames (lor_19 – lor_34). This PUL contains a TonB dependent receptor, along with an experimentally verified rhamnosidase, a β-glucuronyl hydrolase and predicted sulfatases. Interestingly, we also identified in the PUL a new ulvan lyase (LOR_29) which showed no homology to previously reported ulvan lyases making it a founding member of yet another new family of polysaccharide lyases (PL25). Finally, this enzyme prompted us to mine other genomes where we identified additional potential ulvan PULs harboring this gene in other bacterial species. Taken together our report provides further insight into ulvan degradation mechanisms in bacteria and reveals a new family of polysaccharide lyases.

1. Introduction Polysaccharide utilization loci (PUL) refer to large operon structures or clusters of co-regulated genes which code for a set of enzymes involved in polysaccharide breakdown. The paradigm of PUL is the starch utilization loci found in the human gut symbiont, Bacteroides thetaiotaomicron [1]. The typical PUL organization includes genes homologous to susC (a TonB-dependent receptor found in the starch utilization loci) and susD (an oligosaccharide trans-membrane transporter) along with genes encoding a catalog of enzymes implicated in the metabolism of the polysaccharide. The PULs contain genes coding for enzymes involved in polysaccharide degradation (e.g. glycoside hydrolases, polysaccharide lyases), carbohydrate metabolism, enzymes

⁎

1

catalyzing the elimination of the decorations carried on the carbohydrate backbone (e.g. esterases, sulfatases), regulatory genes and genes coding for proteins of unknown function. Because the genes of the PULs are co-regulated, experimental validation of a predicted PUL can be conducted using transcriptomic methods when the bacteria are grown in the presence of selected polysaccharides [2–5]. Currently, predicted and experimentally validated PULs of the human gut Bacteroidetes are listed in the CAZy databank [6], (http://www.cazy.org/PULDB/). PULs have also been predicted in numerous marine microorganisms based on bioinformatics analysis of genomic or metagenomic data [7–11]. However, PUL functions are usually hypothetical and based on sequence homology of glycoside hydrolases present in PULs with known enzymes. This approach allows to suggest a saccharification

Corresponding authors. E-mail addresses: [email protected] (W. Helbert), [email protected] (E. Banin). These authors contributed equally to this work.

http://dx.doi.org/10.1016/j.algal.2017.04.036 Received 12 December 2016; Received in revised form 27 April 2017; Accepted 29 April 2017 2211-9264/ © 2017 Elsevier B.V. All rights reserved.

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moieties: [→ 4)-β-D-GlcA-(1 → 4)-α-L-Rha3S-(1 →], [→ 4)-α-L-IdoA(1 → 4)-α-L-Rha3S(1 →] and [→4)-β-D-Xyl-(1 → 4)-α-L-Rha3S(1 →] [14,15]. Ulva sp. are commonly associated with coastal proliferation due to eutrophicated waters, termed “green tides”, and are grown primarily for supplemental food consumption [16,17]. The world's commercially exploited marine polysaccharides (i.e. alginate, agar and carrageenan) are provided from red and brown algae stock. Despite their high content of carbohydrate - which can reach up to 60% composed of rare residues (Rha, IduA), green algae have not been as much exploited. Interest in Ulva sp. for biofuel feedstock, has recently emerged because these fast growing algae are not competing like other current biofuel biomass sources for land and fresh water [18]. In addition, other biotechnological applications in feed or agronomy, for example, are also envisioned for ulvan. The first gene coding for an ulvan lyase was reported by Collén et al. [19]. The enzyme cleaves the β-(1 → 4)-glycosidic bond between Rha3S and GluA or IduA via β-elimination mechanism. After sequencing the genomes of four ulvanoloytic bacteria [20–22]. Kopel et al. identified LOR_107 (Table 1), a novel ulvan lyase unrelated to the ulvan lyase identified by Collén and colleagues, and as such was the first member of a new family of polysaccharide lyase (PL24 family, ww.cazy.org) [23]. Genes coding for the ulvan-lyase were located in cluster of genes among which contained genes coding for putative rhamnosidases, xylosidases, sulfatases and β-glucuronyl hydrolases. This latter cleaves the glucuronyl residue located at the non-reducing ends of the end-products of ulvan-lyase. Bioinformatic analysis of the Alteromonas sp. LOR genome, led us to identify a PUL encompassing putative β-glucuronyl hydrolase, sulfatase and rhamnosidase potentially involved in the degradation of marine polysaccharide. Here, we report the biochemical characterization of the proteins encoded in a novel ulvan utilization loci, and notably, identify yet another novel ulvan lyase unrelated to all the previously reported enzymes, which is a founding member of a new polysaccharide lyase family (PL25).

Table 1 Genes and protein references cited in this work. Functions, classification in CAZy databank and accession number to Genbank are indicated. Gene locus

Protein name

Function

This work lor_19

LOR_19

lor_20 lor_21

LOR_20 LOR_21

lor_22

LOR_22

lor_23

LOR_23

lor_24 lor_25 lor_26 lor_27 lor_28

LOR_24 LOR_25 LOR_26 LOR_27 LOR_28

Ton-B depended receptor Hypothetical L-Rhamnose mutarotase NAD(P) transhydrogenase NAD(P) transhydrogenase NAD synthetase Hypothetical Sulfatase Rhamnosidase β-Glucuronyl hydrolase New ulvan lyase Sulfatase Hypothetical Hypothetical Sulfatase Rhamnosidase New ulvan lyase New ulvan lyase

lor_29 LOR_29 lor_30 LOR_30 lor_31 LOR_31 lor_32 LOR_32 lor_33 LOR_33 lor_34 LOR_34 nlr_492 NLR_492 plsv_3936 PLSV_3936 Previous work lor_107 LOR_107 nlr_48 NLR_48

Ulvan lyase Ulvan lyase

CAZy

Genbank

WP_032096140.1 WP_032096141.1 WP_032096142.1 WP_032096143.1 WP_032096144.1

GH78 GH105 PL25

GH78 PL25 PL25 PL24 Non classified

WP_032096145.1 WP_032096146.1 WP_032096147.1 WP_032096148.1 WP_032096149.1 WP_052010178.1 WP_032096151.1 WP_032096210.1 – WP_032096152.1 WP_032096153.1 WP_036580476.1 WP_033186995.1 AMA19991.1 AEN28574.1

cascade for the metabolization of putative polysaccharides, which can then be experimentally validated. The first marine PUL to be validated was a PUL which encompassed proteins involved in the degradation of the porphyran, the main cell wall polysaccharide of Porphyra sp., which is often used in the preparation of sushi. Interestingly, this PUL had been transferred laterally from marine Bacteroidetes to Bacteroidetes of the human microbiome [12]. Alginate and laminaran utilization loci were also recently determined experimentally by transcriptomic methods and by characterizing heterologously expressed enzymes [13]. Ulvan is a complex water-soluble sulfated polysaccharide extracted from the cell wall of green algae of the genus Ulvales. It is composed of mainly 3O-sulfate-rhamnose (Rha3S), glucuronic acid (GlcA), iduronic acid (IduA) and xylose (Xyl) distributed in three disaccharide repetition

2. Materials and methods 2.1. Growth conditions of bacterial strains and plasmid constructs Supplementary Tables S1 and S2 list the bacterial strains and primers used in this study. The gene locus, protein name, CAZy classification and Genbank accession number of the various proteins is provided in Table 1. The Native ulvan-degrading isolates were

Fig. 1. Genomic regions in Alteromonas sp. LOR associated with ulvan degradation.

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were inoculated 1:50 (v/v) with transformed cells grown overnight and incubated for ~2.5 h with shaking (150–200 rpm) at 37 °C to reach optical density (595 nm) of 0.6–0.8. Protein expression was induced with 0.3 mM isopropyl 1-thio-β-D-galactopyranoside (IPTG) for 18 h at 16 °C, with shaking. The induced culture was fractionated by centrifugation, forming a pellet. Cells were either lysed chemically (BugBuster protein extraction reagent, EMD Millipore) or by using a French press (cells resuspended in 0.1 M Tris-HCl, 0.2 M NaCl at pH 7.5) followed by centrifugation to remove cell debris. Cell lysate was then purified using either a nickel-charged column (100 mM NiSO4, GE Healthcare) or FPLC (Biorad) followed by ion exchange. After washing, the bound proteins were eluted with a linear gradient of imidazole ranging from 5 to 500 mM. The active fractions were pooled and protein purification was verified using SDS polyacrylamide gel electrophoresis stained with instant blue (Expedeon). 2.3. Activity assays and degradation product analysis The ulvan substrate used for activity assays and degradation kinetics was provided by CEVA (Pleubian, France). The polysaccharide was extracted from Ulva rotundata and purified as described by Collén et al. [19]. Cloned enzymes were initially screened for ulvanolytic activity using the ulvan polysaccharide or ulvan oligomers as substrate by measuring the formation (UL activity) or breaking (β-glucuronyl hydrolase) of double bonds at 235 nm [24]. The assay solution was composed of 1 g/l ulvan in 0.1 M Tris-HCl (Sigma), 0.2 M NaCl at pH 7.5 and with either purified proteins (5 μl of 160 μg/ml protein solution) or cell lysate (5 μl) in 180 μl total volume unless otherwise specified. For cell lysate preparation batch cultures were grown overnight and protein expression inducted as described in the previous section followed by cell lysis using PopCulture reagent (Novagen) according to manufacturer's instructions. The spectrophotometer (Synergy™ 4 Hybrid Microplate Reader, BioTek) was set to read at intervals and the change in absorbance at 235 nm was monitored. The optimal temperature and pH were determined using the initial velocity which corresponds to the linear slope recorded by spectroscopy at 235 nm. For the pH assays, 0.1 g/l ulvan was dissolved in buffers with a pH range of 6–9. Each buffer was composed of 0.1 M Tris (Sigma), 0.2 M NaCl adjusted with HCl to the tested pH. Similarly, the optimal temperature was determined by using a range of temperature from 20 °C to50°C, in 10 °C increments. Briefly, 0.1 g/l ulvan was dissolved in the optimal buffer found in this study (pH 7.5). 5 μl of purified enzyme (160 μg/ml) was added directly to a 180 μl prewarmed ulvan solution and the increase in absorbance at 235 nm were followed in a spectrophotometer (Synergy™ 4 Hybrid Microplate Reader, BioTek) set to read at 1 min intervals at the appropriate temperature. The p-nitrophenol (PNP) colorimetric assay was used to assess enzyme cleavage of bonds that liberated specific sugars. Increased absorbance was measured at 415 nm, indicating that PNP is released from its synthetic substrate (i.e. sulfate, rhamnose, and xylose).

Fig. 2. Enzymatic activity of LOR_34, LOR_28 and LOR_29. (A) 4-Nitrophenyl- α-Lrhamnose incubated with cell lysate derived from over expressed LOR_34 was followed by UV415nm. Rhamnosidase activity is indicated by the increase in absorbance. (B) Ulvan modified by LOR_29 incubated with cell lysate derived from over expressed LOR_28 was followed by UV235nm absorbance. Glycoside hydrolase activity is indicated by the decrease in UV235nm absorbance. (C) Ulvan incubated with cell lysate derived from over expressed LOR_29 was followed by UV235nm absorbance. Ulvan lyase activity is indicated by the increase in absorbance. In all experiments cell lysate with an empty vector, pET28, served as control. The results presented are average of three independent experiments carried out in triplicates, errors bars indicate standard deviation.

cultured in Marine Broth (Difco) at 25–28 °C, while Escherichia coli strains were grown in Luria Bertani (LB medium, Difco) at 37 °C. Where indicated (Supplementary Table S1), antibiotics were added to culture media. When primers were designed for the genes of interest from genomic DNA of listed native marine isolates, a His-tag was added on the C-terminus and primers and genes were amplified with (e.g., LOR_29) and without their native signal peptide (e.g., LOR_107d) (Supplementary Table S2). Both the gene insert and expression vector were digested using relevant restriction enzymes (details in Supplementary Table S2), purified and then ligated into an expression plasmid. T7 Express competent E. coli (NEB) were then transformed with recombinant plasmids (Supplementary Table S1).

2.4. Size exclusion chromatography – analytical/semi-preparative gel permeation chromatography Degradation kinetics on ulvan was measured after various incubation periods with purified ULs. Oligo-ulvan end products were then purified and analyzed by 1H NMR. 100 g/l ulvan in 0.1 M Tris-HCl, 0.2 M NaCl at pH 7.5 was incubated with 60 μg/mL purified enzymes at room temperature for 24 h. Ulvan degradation kinetics were monitored by analytical gel permeation chromatography. Samples (20 μL) were injected on a Superdex 200 (10/300 GL, GE Healthcare) and a peptide HR (10/300 GL, GE Healthcare) column coupled in series. Elution was conducted with an Ultimate 3000 HPLC system (ThermoScientific, Dionex) operating at 0.5 ml/min with 50 mM (NH4)2CO3 (pH 8) as the eluent. Detection of products was achieved using a RSLC UV

2.2. Protein expression and purification Overexpression of PUL enzymes was performed using transformed E. coli strains (listed in Supplementary Table S1), followed by cell lysis and purification of His-tagged proteins. Batch cultures (50–500 mL) 41

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(caption on next page)

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Fig. 3. Degradation pathway of ulvan. Ulvan is composed mainly of the repetition of sulfated rhamnose (R3S) linked either to glucuronic (GlcA) or iduronic residues (IduA). The unsaturated residue (4-deoxy-L-threo-hex-4-enopyranosiduronic acid, Δ) linked to the sulfated rhamnose occurs at the non-reducing end and is exposed after cleavage of the glycosidic linkage by ulvan lyase. When it is cleaved by the β-glucuronyl hydrolase (GH_105), the cyclic form spontaneously rearranges to a linear 4-deoxy-1-threo-5-hexosulose urinate.

2.4

charides at 0.5% (w/v)) were injected on three Superdex 30 columns (GE Healthcare, 2.6 × 60 cm) mounted in series. Elution was conducted at a flow rate of 1.5 ml/min. Oligosaccharides were collected with a Foxy R1 collector (Teledyne ISCO) controlled by AZUR software (Datalys).

-R3S

2.2

-R3S-Xyl-R3S

2.0 Refractive index (R.I.)

1.8

2.5. NMR spectroscopy

1.6 T= 24 h 1.4 1.2 1.0

1

H NMR spectra were recorded at 353 K on a 400 MHz Avance DRX400 spectrometer (Bruker) in deuterated water as solvent, and calibrated against the residual signal of the solvent. Prior to analysis, samples were exchanged two times in D2O and lyophilized, then were re-dissolved in D2O (99.97 atom % D). Chemical shifts are expressed in ppm in reference to an external standard (trimethylsilylpropionic acid). The HOD signal was not suppressed.

T= 6 h

T= 30 min

0.8 0.6 T= 5 min 0.4 0.2

2.6. Statistical analysis

T= 0

All enzymatic assays described and presented in the work were performed in three independent experiments in triplicates. Error bars represent standard deviation which was calculated using the statistical model in GraphPad Prism, version 5.0.

0.0 10.0

15.0

20.0

25.0

30.0

35.0

Retention time (min.) Fig. 4. Degradation kinetics of ulvan incubated with LOR_29. Gel permeation chromatography shows that the ulvan polysaccharide is rapidly converted into oligo-ulvan products. The most abundant oligomers are disaccharides (Δ-R3S), followed by tetra saccharides (Δ-R3S-Xyl-R3S). The experiment was repeated three times in triplicates. The results are the representation of one experiment.

3. Results and discussion

detector (ThermoScientific, Dionex) operating at 235 nm, along with an IOTA 2 refractive index detector (Precision Instrument). Oligo-ulvan end-products were purified using filtered digested ulvan (0.22 μm nylon clarinet syringe filters, Agela technology). Samples (mL of oligosac-

Mining the genome of Alteromonas sp. LOR revealed three clusters of genes containing sulfatases, glycoside hydrolases and polysaccharide lyases (Fig. 1). Two clusters contained the genes coding for the previously reported ulvan lyases belonging to the PL24 family [23]. These genes are located adjacent to genes coding for predicted

3.1. Polysaccharide degrading enzymes of Alteromonas sp. LOR are clustered in the genome

Fig. 5. H-NMR spectra of the two purified dominant LOR_29 end-products. (A) Spectrum of the purified tetrasaccharide (DP4) Δ-R3S-Xyl-R3Sα/β. (B) Spectrum of the purified disaccharide (DP2) Δ-R3Sα/β. The experiment was repeated three times in triplicates. The results are the representation of one experiment.

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LOR_34 with the products obtained after degradation with ulvan lyase and β-glucuronyl hydrolase (i.e. the trisaccharide: β-α-L-Rha3S-(1 → 4)α-L-IdoA/β-D-GlcA-(1 → 4)-α-L-Rha3S), showed no degradation (as measured by chromatography). This suggests that LOR_34 is active on neutral rhamnose and that desulfation of the oligosaccharide must be achieved before cleavage of rhamnose. LOR_28 is classified with the GH105 family, which encompasses both α- and β-glucuronyl hydrolases [25]. LOR_28 shows only 37% sequence identity with the β-glucuronyl hydrolase of Nonlabens ulvanivorans known to be active on oligo-ulvans. Incubation of ulvan-lyase (LOR_107, PL24) end-products with LOR_28 was monitored by spectrophotometry and a rapid decrease of the absorbance at 235 nm demonstrating that LOR_28 was a β-glucuronyl hydrolase active on oligo-ulvans (Fig. 2B). Three sulfatases are predicted in the PUL (LOR_26, LOR_30, and LOR_33). They all belong to the formyl-glycine dependent sulfatases which require the post-translational modification of cysteine or serine to the catalytic formylglycine amino acid. LOR_26, and LOR_30 were obtained soluble but sulfatase activity could not be confirmed. Ulvan, oligo-ulvans and the synthetic substrate PNP-sulfate were used but desulfation was never detected. The lack of activity could be explained by the absence of post-translational modification of the catalytic amino acids in E. coli. The proteins with unknown function LOR_29 and LOR_31 were obtained soluble. LOR_31 was screened on ulvan, oligo-ulvans and a set of PNP-glycosides without revealing any potential activity on ulvan. However, when LOR_29 was incubated with ulvan, we observed by spectrometry a rapid increase of the absorbance at 235 nm synonymous to the formation of unsaturation at the non-reducing ends of oligosaccharides (Fig. 2C). Altogether, these observations suggested that the PUL investigated was directed toward ulvan degradation and that LOR_29 was the first member of a novel family of polysaccharide lyase PL25 (B. Henrissat, personal communication). Taken together these results allow us to suggest a degradation pathway of ulvan starting with the novel ulvan lyase (LOR_29) followed by the action of β-glucuronyl hydrolase (LOR_28), sulfatases (LOR_26, 30 and 33) and ultimately by rhamnosidases (LOR_27 and 34) (Fig. 3).

Fig. 6. Comparison of ulvan degradation products by purified heterologous overexpressed ulvan lyases. Gel permeation chromatography shows a comparison of endproducts obtained after incubating the tetrasaccharide mixture (DP4) of Δ-R3S-IduA-R3S, Δ-R3S-GluA-R3S and Δ-R3S-Xyl-R3S with the ulvan lyases LOR_107, LOR_29, or its homolog NLR_492. The DP2 (Δ-R3S) and DP4 spectra represents purified oligo-ulvan isolates without enzyme incubation. The experiment was repeated three times in triplicates. The results are the representation of one experiment.

3.3. Biochemical characterization of the novel ulvan lyase LOR_29 The protein sequence of LOR_29 did not reveal homology to the previously reported ulvan lyases, LOR_107 (PL24) [23] or NLR_48 (not classified) [19]. Instead, protein sequences homologous to LOR_29 were found in two recently sequenced genomes Pseudoalteromonas sp. PLSV (PLSV_3936, 76% identity) and N. ulvanivorans (NLR_492, 53% identity), as well as Paraglaciecola agarilytica (76% identity), Paraglaciecola chathamensis (75% identity), Catenovulum agarivorans (61% identity) and Pseudoalteromonas haloplanktis (57% identity), among others. This suggests that LOR_29 is the first representative of a new family of polysaccharide lyases. LOR_29 and NLR_492 were overexpressed soluble and their molecular mass were observed at 52 kDa for LOR_29 and 55 kDa for NLR_492, as expected (Supplementary Fig. S1). The biochemical parameters of LOR_29 were determined by measuring the initial velocity with spectrometry at 235 nm. The optimum pH was measured as 7.5 and the optimum temperature was 45 °C (Supplementary Fig. S2). Degradation kinetics of LOR_29 was monitored by chromatography revealing its endo-acting mode of action (Fig. 4). The chromatogram corresponding to undigested ulvan showed a single peak eluting in the exclusion volume at about 15 min. As the degradation of the substrate proceeded, the signal of the polymer fraction became broader and shifted toward longer elution, indicating a decrease in the molecular mass associated to an increase of the polydispersity. After 24 h incubation, the elution time observed for the end-products suggested the main product was a disaccharide (31 min), and in lower amounts a

glucuronyl hydrolases (GH105, GH88), rhamnosidases (GH78) and xylosidase (GH43). Because these clusters lack TonB-dependent receptors and a transmembrane carbohydrate transporter, gene expression may be regulated differently than the emblematic PUL. The last cluster of genes (lor_19 through lor_34, corresponding identifiers can be found in Table 1) contained a TonB dependent receptor, along with predicted sulfatases (lor_26, lor_30 and lor_33), glucuronyl hydrolase (GH 105: lor_28) and rhamnosidases (GH 78: lor_27 and lor_34). This cluster had the characteristic of a PUL targeting sulfated polysaccharides composed of rhamnose. Interestingly, the presence of a gene coding for glucuronyl hydrolase (GH105, lor_28) suggested that a polysaccharide lyase could also be present in the PUL since glucuronyl hydrolase cleaves the endproducts of lyases, yet no homolog to the known ulvan lyases was identified. Thus, we have investigated the catalytic functions of the proteins coded by this PUL.

3.2. Cloning, overexpression and activity assessment The genes (lor26 to lor_34) were cloned and coded proteins were overexpressed in E. coli. The predicted GH78 (LOR_34) and GH105 (LOR_28) were obtained soluble and their catalytic function was validated experimentally. The LOR_34 cleaved PNP-α-L-rhamnoside confirming the predicted rhamnosidase activity (Fig. 2A). Incubating 44

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Fig. 7. PULs encompassing gene homologs to ORF_29 encoding for the novel ulvan lyase. The PULs or cluster of gene suggest potential ulvan degradation activity of the corresponding strains.

the GlcA residue [23]. Interestingly, NLR_492 showed the same modality of degradation as LOR_29 suggesting that the catalytic properties of the ulvan-lyase of this novel family were conserved.

tetrasaccharide (28 min). The end-products were purified using preparative size exclusion chromatography. Based on previously reported oligo-ulvans 1H NMR spectra [19], the disaccharide was identified (Fig. 5) as a glucuronyl residue-linked sulfated rhamnose (Δ-R3S). The tetrasaccharide did not contain uronic residues and had the Δ-R3S-Xyl-R3S structure. These results showed that both iduronic and glucuronic acid residues were cleaved by the enzymes and suggested, also that tetrasaccharides containing uronic residues are cleaved. When comparing cleavage preference of the tetrasaccharide mixture of Δ-R3S-IduA-R3S, Δ-R3S-GluA-R3S and Δ-R3S-Xyl-R3S between LOR_29 and LOR_107 (PL24 family), we observed that the tetrasaccharide was completely converted to di-saccharides by LOR_29, with undigested Δ-R3S-Xyl-R3S fraction eluting at 28 min (Fig. 6). This differs from the tetrasaccharides incubated with LOR_107, which yielded a low conversion to disaccharides due to its specificity toward

3.4. Mining other genomes for Alteromonas sp. LOR_29 homologs and identification of additional ulvan utilization loci Searching bacterial genomes in Genbank and RAST [26] revealed that genes encoding for homologous protein to LOR_29 were located in PULs or in gene clusters (Fig. 7). The PULs encompassing LOR_29 was conserved in Pseudoalteromonas PLSV [20] and Paraglaciecola agarilytica NO2 [27]. A long PUL containing a LOR_29 homolog to with a different gene organization compared to the other clusters, was found in a sequenced contig of Erythrobacter marinus [28]. This PUL encodes all the enzymes required for complete ulvan degradation such as βglucuronyl hydrolase (GH105, GH88), rhamnosidase (GH78), xylosi45

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dase (GH3) and several sulfatases. Two TonB dependent receptors were present and associated to several ABC transporters. In Catenovulum agarivorans DS-2 [29], a homolog to LOR_29 occurred with another ulvan lyase belonging to PL24 family. Along with ulvan lyase and the TonB dependent receptor, only one predicted rhamnosidase was observed. This suggested that the other hypothetical proteins found in the PUL were probably involved in ulvan degradation and/or other part of the PUL is located in another place in the genome. The first gene of ulvan lyase was described in N. ulvanivorans, and this gene was found in a PUL. Interestingly, N. ulvanivorans genome encoded for another ulvanolytic system made of NLR_492 (homolog of LOR_29), rhamnosidases of GH106 family and sulfatases. 4. Conclusion The current study utilized a bioinformatics approach to mine the genome of an ulvan degrading bacterial strain Alteromonas sp. LOR. Using specific criteria, such as the presence of TonB-dependent receptors, glucuronyl hydrolases, rhamnosidases and sulfatases, we were able to identify a novel PUL involved in ulvan degradation. The PUL contains a functional β-glucuronyl hydrolase and rhamnosidase, as well as several predicted sulfatases. Surprisingly, no homolog to the previously identified ulvan lyases was found in the cluster. This led us to test various hypothetical proteins within the PUL and we were able to identify a new ulvan lyase, which is the first representative of a new family of polysaccharide lyases PL25 (personal communication Bernard Henrissat, CAZy). Furthermore, using this enzyme as a basis to mine the database we identified additional putative ulvan related PULs in other bacterial species. This study not only promotes our understanding of the complex mechanisms associated with microbial degradation of ulvan but also supported the identification of yet another new family of polysaccharide lyases that is fairly wide spread in bacteria. Acknowledgements We thank Claire Boisset and Laurine Buon from CERMAV for their excellent chromatography services and Michael Abeles for his support. This research was partially funded by an Infrastructure Grant from the Israel Ministry of Science, Technology and Space (H50/2013) awarded to EB. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.algal.2017.04.036. References [1] K.L. Anderson, A.A. Salyers, Genetic evidence that outer membrane binding of starch is required for starch utilization by Bacteroides thetaiotaomicron, J. Bacteriol. 171 (1989) 3199–3204. [2] E.C. Martens, H.C. Chiang, J.I. Gordon, Mucosal glycan foraging enhances fitness and transmission of a saccharolytic human gut bacterial symbiont, Cell Host Microbe 4 (2008) 447–457. [3] E.C. Martens, E.C. Lowe, H. Chiang, N.A. Pudlo, M. Wu, N.P. McNulty, D.W. Abbott, B. Henrissat, H.J. Gilbert, D.N. Bolam, J.I. Gordon, Recognition and degradation of plant cell wall polysaccharides by two human gut symbionts, PLoS Biol. 9 (2011), http://dx.doi.org/10.1371/journal.pbio.1001221. [4] N.P. McNulty, M. Wu, A.R. Erickson, C. Pan, B.K. Erickson, E.C. Martens, N.A. Pudlo, B.D. Muegge, B. Henrissat, R.L. Hettich, J.I. Gordon, Effects of diet on resource utilization by a model human gut microbiota containing Bacteroides cellulosilyticus WH2, a symbiont with an extensive glycobiome, PLoS Biol. 11 (2013), http://dx.doi.org/10.1371/journal.pbio.1001637. [5] J. Despres, E. Forano, P. Lepercq, S. Comtet-Marre, G. Jubelin, C.J. Yeoman, M.E.B. Miller, C.J. Fields, N. Terrapon, C. Le Bourvellec, C. Renard, B. Henrissat, B.A. White, P. Mosoni, Unraveling the pectinolytic function of Bacteroides xylanisolvens using a RNA-seq approach and mutagenesis, BMC Genomics 17 (2016), http://dx.doi.org/10.1186/s12864-016-2472-1. [6] N. Terrapon, V. Lombard, H.J. Gilbert, B. Henrissat, Automatic prediction of polysaccharide utilization loci in Bacteroidetes species, Bioinformatics 31 (2014) 647–655.

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