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In vitro and in vivo characterization of three Cellvibrio japonicus glycoside hydrolase family 5 members… Attia, Mohamed A; Nelson, Cassandra E; Offen, Wendy A; Jain, Namrata; Davies, Gideon J; Gardner, Jeffrey G; Brumer, Harry Feb 17, 2018

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Attia et al. Biotechnol Biofuels  (2018) 11:45  https://doi.org/10.1186/s13068-018-1039-6RESEARCHIn vitro and in vivo characterization of three Cellvibrio japonicus glycoside hydrolase family 5 members reveals potent xyloglucan backbone-cleaving functionsMohamed A. Attia1,2, Cassandra E. Nelson3, Wendy A. Offen4, Namrata Jain1,2, Gideon J. Davies4, Jeffrey G. Gardner3 and Harry Brumer1,2,5,6*Abstract Background: Xyloglucan (XyG) is a ubiquitous and fundamental polysaccharide of plant cell walls. Due to its structural complexity, XyG requires a combination of backbone-cleaving and sidechain-debranching enzymes for complete deconstruction into its component monosaccharides. The soil saprophyte Cellvibrio japonicus has emerged as a genetically tractable model system to study biomass saccharification, in part due to its innate capacity to utilize a wide range of plant polysaccharides for growth. Whereas the downstream debranching enzymes of the xyloglucan utilization system of C. japonicus have been functionally characterized, the requisite backbone-cleaving endo-xyloglu-canases were unresolved.Results: Combined bioinformatic and transcriptomic analyses implicated three glycoside hydrolase family 5 subfam-ily 4 (GH5_4) members, with distinct modular organization, as potential keystone endo-xyloglucanases in C. japoni-cus. Detailed biochemical and enzymatic characterization of the GH5_4 modules of all three recombinant proteins confirmed particularly high specificities for the XyG polysaccharide versus a panel of other cell wall glycans, including mixed-linkage beta-glucan and cellulose. Moreover, product analysis demonstrated that all three enzymes generated XyG oligosaccharides required for subsequent saccharification by known exo-glycosidases. Crystallographic analysis of GH5D, which was the only GH5_4 member specifically and highly upregulated during growth on XyG, in free, product-complex, and active-site affinity-labelled forms revealed the molecular basis for the exquisite XyG specific-ity among these GH5_4 enzymes. Strikingly, exhaustive reverse-genetic analysis of all three GH5_4 members and a previously biochemically characterized GH74 member failed to reveal a growth defect, thereby indicating functional compensation in vivo, both among members of this cohort and by other, yet unidentified, xyloglucanases in C. japoni-cus. Our systems-based analysis indicates distinct substrate-sensing (GH74, GH5E, GH5F) and attack-mounting (GH5D) functions for the endo-xyloglucanases characterized here.Conclusions: Through a multi-faceted, molecular systems-based approach, this study provides a new insight into the saccharification pathway of xyloglucan utilization system of C. japonicus. The detailed structural–functional charac-terization of three distinct GH5_4 endo-xyloglucanases will inform future bioinformatic predictions across species, and provides new CAZymes with defined specificity that may be harnessed in industrial and other biotechnological applications.Keywords: Xyloglucan, Saccharification, Glycoside hydrolase, Cellvibrio japonicus, Saprophyte© The Author(s) 2018. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creat iveco mmons .org/licen ses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creat iveco mmons .org/publi cdoma in/zero/1.0/) applies to the data made available in this article, unless otherwise stated.Open AccessBiotechnology for Biofuels*Correspondence:  brumer@msl.ubc.ca 1 Michael Smith Laboratories, University of British Columbia, 2185 East Mall, Vancouver, BC V6T 1Z4, CanadaFull list of author information is available at the end of the articlePage 2 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 BackgroundRenewable plant biomass is envisioned as a promis-ing alternative to fossil petroleum for the production of liquid fuels and high-value chemicals [1, 2]. Plant cell walls are, however, chemically and structurally complex in nature and require harsh thermo-chemical treatment to yield fermentable sugars. Such processes often gen-erate undesirable by-products that inhibit subsequent microbial conversion [3]. In light of their ability to cata-lyze the degradation of recalcitrant plant cell walls under ambient conditions, enzymes from saprophytic micro-organisms constitute an attractive palette of biocatalysts for improved biomass saccharification [4]. The discovery and characterization of new enzymes from saprophytes is thus central to advancing biotechnology and, not least, underpins fundamental understanding of the biological roles of these micro-organisms in the global carbon cycle.The Gram-negative bacterium, Cellvibrio japonicus Ueda107 (formerly, Pseudomonas fluorescens subsp. cel-lulosa) has emerged as a model saprophytic micro-organ-ism with a demonstrated ability to utilize nearly all plant cell wall polysaccharides, including cellulose, xylans, mannans, arabinans, and pectins [5, 6]. Indeed, sequenc-ing of the C. japonicus genome in 2008 revealed vast array of carbohydrate-active enzymes (CAZymes [7]) predicted to be involved in plant cell wall saccharifica-tion [8]. The recent development of genome editing tech-niques for C. japonicus has further advanced the biology and bioengineering of this bacterium in biomass conver-sion [9–13].The xyloglucans (XyG) comprise an important fam-ily of cell wall matrix polysaccharides, which are ubiqui-tous and abundant across the plant kingdom [14, 15]. In dicots, XyGs may constitute up to 25% of the primary cell wall dry-weight, with lower amounts found in conifers (10%) and grasses (< 5%) [16, 17]. Structurally, XyGs have brush-like architectures built upon a linear, cellulosic β(1→4)-d-glucan backbone that is extensively branched with α(1→6)-xylopyranosyl residues at regular intervals. Further elaboration of these branch points with diverse monosaccharides and acetyl groups is dependent on the species and tissue of origin [18, 19]; presently ca. 20 dis-tinct sidechain saccharide compositions are known [20, 21]. The structure of the canonical dicot (fucogalacto)xyloglucan is shown in Fig.  1a. Due to this structural complexity, complete XyG saccharification requires the concerted action of numerous backbone-cleaving endo-xyloglucanases and side-chain-cleaving exo-glycosidases [22, 23].As part of our ongoing effort to elucidate the xyloglucan (XyG) utilization system of C. japonicus, we functionally characterized a multi-gene XyG utilization locus (XyGUL) in the C. japonicus genome via a combination of genetics, enzymology, and structural biology. This XyGUL encodes the three exo-glycosidases required for (fucogalacto)xylo-glucan sidechain cleavage (a GH95 α-l-fucosidase, a GH35 β-galactosidase, and a GH31 α-xylosidase) together with a predicted TonB-dependent transporter (TBDT) (Fig. 1b, c) [13, 24, 25]. A highly specific β-glucosidase, Bgl3D, which is encoded elsewhere in the genome, works in concert with the exo-glycosidases of the XyGUL to effect the complete saccharification of XyG oligosaccharides (XyGOs) in the periplasm (Fig. 1b, c) [26]. Noting that this locus likewise lacked an associated endo-xyloglucanase, we also provided biochemical and structural evidence that the lone, secreted C. japonicus GH74 member (Fig.  1b, c) could efficiently generate the  Glc4-based XyGOs required by the down-stream exo-glycosidases [27].KeyGlcpGalpXylpL-Fucpa=0, b=0, c=0: XXXGa=0  b=1, c=0: XXLGa=0, b=1, c=1: XXFGa=1, b=0, c=0: XLXGa=1, b=1, c=0: XLLGa=1, b=1, c=1: XLFGGH31GH35TBDTGH95GH74CJA_2706 0172_AJC7742_AJCGH5FCJA_2959GH5DCJA_3010GH5ECJA_3337ab(reducing end)nXXXG- type XyGa bcCjGH5DTonB-ExbBDComplexTBDTCjGH74 CjGH5ECjGH5FAfc95ABgl35AXyl31ABgl3DcBgl3DCJA_1140Fig. 1 Xyloglucan (XyG) and the xyloglucan utilization system in C. japonicus. a Structure of dicot XXXG-type fucogalacto-XyG. XyG substructure nomenclature is according to [20]. b C. japonicus genes involved in XyG utilization. Genes encoding backbone-cleaving endo-xyloglucanases (GH5 and GH74) are indicated in navy blue, genes encoding side-chain-cleaving exo-glycosidases (GH35 β-galactosidases; GH31 α-xylosidases and GH95 α-l-fucosidase) are in cyan, and the TonB dependent transporter (TBDT) is shown in green. c Spatial model of XyG utilization in C. japonicus Page 3 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 As we now show, genetic deletion of this GH74 endo-xyloglucanase did not, however, impede the growth of C. japonicus on the polysaccharide, which suggested the involvement of additional, unidentified endo-xyloglu-canases. Hence, we also explored the in vitro and in vivo function of three candidate endo-xyloglucanases from GH5 subfamily 4 (GH5_4) [28], guided by bioinformatic and transcriptomic analyses. Utilizing a combination of reverse genetics, enzymology, and structural biology, the present study provides a new insight into the upstream deconstruction of XyG.Results and discussionTranscriptomic analysis reveals a potential keystone endo‑xyloglucanase from Glycoside Hydrolase (GH) family 5, subfamily 4We previously showed via quantitative PCR (qPCR) that the C. japonicus gene cluster containing xyl31A (CJA_2706), bgl35A (CJA_2707), CJA_2709, and afc95A (CJA_2710) (Fig. 1b), was up-regulated during growth on xyloglucan-containing medium [13]. Biochemical char-acterization confirmed that xyl31A, bgl35A, and afc95A encode a XyGO-specific GH31 α-xylosidase, GH35 β-galactosidase, and GH95 α-l-fucosidase, respectively, while CJA_2709 was predicted to encode a TonB-depend-ent transporter (TBDT) [13, 24, 25]. To aid identification of potential C. japonicus endo-xyloglucanases acting upstream of these enzymes, a comprehensive expression analysis via RNAseq was performed in the present study. Samples were collected from both exponentially grow-ing and stationary phase cells grown on glucose or xylo-glucan as the sole carbon source to allow for analyses of gene expression based on early-stage substrate detection (Additional file  1: Figure S1), late-stage substrate detec-tion (Additional file 1: Figure S2A), or growth rate (Addi-tional file 1: Figure S2B).During exponential growth, there were 27 CAZyme-encoding genes significantly up-regulated on XyG, including the four genes of the C. japonicus XyG cluster, which corroborated previous qPCR results (Additional file  1: Table  S1). Notably, CJA_3010, which encodes a GH5 subfamily 4 (GH5_4) member previously anno-tated as cel5D, was the highest upregulated gene fol-lowed by the XyG cluster genes CJA_2709 (encoding a predicted TBDT) and CJA_2706 (xyl31A, encoding a GH31 α-xylosidase) [8]. Among the large and func-tionally diverse GH5 family [28], subfamily 4 is the only subfamily known to contain predominant endo-xylo-glucanases [23], which suggested a keystone role for this enzyme in xyloglucan utilization by C. japonicus. Notably, CJA_2477 (previously annotated as gly74 [8]; Fig. 1b) was not significantly up-regulated during growth on XyG, despite the encoded GH74 endo-xyloglucanase being previously shown to have high, specific activity for this polysaccharide [27]. Instead, CJA_2477 appeared to be constitutively expressed at a low level (RPKM levels in the 100–200 range), as were 14 other predicted CAZyme-encoding genes (Additional file 1: Table S2).The remaining CAZyme genes up-regulated during exponential growth are predicted to have roles in the degradation of a diverse set of polysaccharides, which suggests that there is complex cross-regulation of expres-sion. As xyloglucan is unlikely to be encountered alone during the saprophytic growth habit of C. japonicus, these results are suggestive of xyloglucan degradation being one component of a sophisticated plant cell wall degradation response. Congruently, when comparing the stationary phase C. japonicus cells growing on XyG and glucose, only two genes of the XyG cluster, bgl35A and afc95A were still up-regulated on XyG, together with 33 additional predicted and confirmed hemicellulase- and pectinase-encoding genes (Additional file 1: Figure S2A, Table S3). Additionally, when comparing the exponential phase to the stationary phase for xyloglucan-grown cells, we observed that there was a growth-phase-dependent response manifested as a significant shift in the suite of expressed CAZyme genes (Additional file 1: Figure S2B, Table  S4). Specifically, these differentially expressed CAZyme genes were not predicted to be XyG-specific, based on their CAZy family membership, which sug-gested they are part of regulatory circuit that responds generally to polysaccharides. Similar growth-phase-dependent responses have been previously observed during cellulose utilization by C. japonicus [12] and Clostridium thermocellum (now Ruminiclostridium ther-mocellum) [29, 30].Bioinformatic analysis and recombinant production of GH5_4 members from C. japonicusSpurred  on by the implication of the GH5 subfamily 4 (GH5_4) member encoded by CJA_3010 in xyloglucan utilization by C. japonicus, we searched the genome for potential homologs. C. japonicus encodes 15 GH5 mem-bers, of which only three belong to subfamily 4 ([8] see http://www.cazy.org/b776.html): the aforementioned CJA_3010 (GenBank ACE84905.1, previously annotated as cel5D [8]), CJA_3337 (GenBank ACE83841.1, previ-ously annotated as cel5E [8]), and CJA_2959 (GenBank ACE86198.1, previously annotated as cel5F [8]). Pro-tein sequence analysis revealed that each of these gene products had a unique, multi-modular architecture that suggested the possibility of distinct cellular localiza-tion and biological function (Additional file  1: Figure S3). Considering the lack of demonstrable activity on cellulose and high activity on xyloglucan (vide infra), Page 4 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 the corresponding encoded enzymes are referred to as CjGH5D, CjGH5E, and CjGH5F hereafter.The highly up-regulated CJA_3010 encodes a signal peptidase II lipoprotein signal peptide (predicted by LipoP 1.0 [31]), followed by a serine-rich linker and a GH5_4 catalytic module, and was thus predicted to be anchored extracellularly in the outer membrane by N-ter-minal cysteine lipidation (Additional file  1: Figure S3). CJA_3337 encodes an N-terminal signal peptide (pre-dicted by SignalP 4.0 [32]) and two carbohydrate-binding modules (CBMs [33]), CBM2 and CBM10, in train with a GH5_4 catalytic module (Additional file 1: Figure S3). CJA_2959 encodes a signal peptide (predicted by SignalP 4.0 [32] a Fibronectin type III (FN3) domain, an unde-fined region, and a C-terminal GH5_4 catalytic module (Additional file 1: Figure S3). The presence of signal pep-tides, and CBMs in the case of CjGH5E, is indicative of extracellular secretion of both CjGH5E and CjGH5F.Amino acid alignment of the catalytic modules of CjGH5D, CjGH5E, and CjGH5F demonstrate conserva-tion of the two catalytic glutamate residues, but low to moderate overall sequence conservation (26–45% iden-tity). Notably, alignment with endo-xyloglucanases from Bacteroides ovatus [22], Paenibacillus pabuli [34], and a rumen metagenome [35] suggests that the C. japonicus proteins are members of subfamily 4 (Additional file  1: Figure S4, Table  S5). GH5_4 is one of the largest GH5 subfamilies and contains, in addition to specific endo-xyloglucanases, promiscuous endo-β(1,4)-glucanases, strict cellulases, and mixed-linkage endo-β(1,3)/β(1,4)glucanases (reviewed in [23, 28]). As such, we undertook the recombinant production and enzymological charac-terization of the three C. japonicus GH5_4 members to precisely define their catalytic activities in the context of potential biological function.Our initial attempts to produce the full-length, multi-modular proteins recombinantly in E. coli by replace-ment of the native signal peptides with an N-terminal hexahistidine  (His6) purification tag were consistently unsuccessful: intact protein mass spectrometry revealed proteolytic instability of  His6-SRL-GH5D, while  His6-CBM2-CBM10-GH5E and  His6-FN3-GH5F had very poor production yields (data not shown). In con-trast,  His6-GH5D (Additional file  1: Figure S3A) was produced as a stable, intact, active protein (calculated mass, 44,222.2 Da; observed by ESI–MS, 44,222.6 Da) in excellent yield (150  mg  L−1). Likewise, our attempts to produce the individual catalytic modules of CjGH5E and CjGH5F as N-terminally  His6-tagged constructs (Addi-tional file 1: Figure S3B, C) were successful  (His6-CjGH5E calculated mass, 41,367.1  Da; observed by ESI–MS, 41,370.1 Da,  His6-CjGH5F calculated mass, 40,253.8 Da; observed by ESI–MS 40,253.9 Da) with approximate pro-duction yields of 14 and 9 mg L−1, respectively.CjGH5_4 enzymes are highly efficient, specific endo‑xyloglucanasesInformed by the subfamily membership of the three GH5_4 members, we anticipated that these enzymes might exhibit significant endo-hydrolytic activity towards XyG. Hence, this polysaccharide was used to determine pH and temperature optima. CjGH5D, CjGH5E, and CjGH5F each exhibited approximately bell-shaped pH profiles, with the highest activity achieved in 50  mM phosphate buffer (pH 7.5 in the  case of CjGH5D and CjGH5E, and pH 7.0 in the case of CjGH5F; Additional file  1: Figure S5). When the three enzymes were incu-bated with XyG at different temperatures over the course of 10 min, the optimum temperatures were identified as 50 °C (CjGH5D and CjGH5F) and 55 °C (CjGH5E) (Addi-tional file 1: Figure S5). To determine substrate specificity of the three GH5_4 members, a panel of nine soluble pol-ysaccharide substrates were screened under these opti-mal conditions. Indeed CjGH5D, CjGH5E, and CjGH5F all displayed high specific activity toward XyG (Table 1). No detectable activity toward barley mixed-linkage Table 1 Activity of CjGH5_4 enzymes against different polysaccharide substratesAssays conducted at pH 7.5 (CjGH5D and CjGH5E) or pH 7 (CjGH5F). Recombinant enzymes were incubated at 50 °C (CjGH5D and CjGH5F) or 55 °C (CjGH5E) with the different tested substratesND not determined due to poor specific activityEnzyme catalytic domains Substrate Km mg mL−1 kcat  s−1 Specific activity µmol (min mg)−1CjGH5D XyG < 0.025 30.3 ± 0.4 43.3 ± 1.9CjGH5E XyG 0.020 ± 0.002 10.3 ± 0.1 15.1 ± 0.1Hydroxyethylcellulose (HEC) ND ND 0.070 ± 0.004Carboxymethylcellulose (CMC) ND ND 0.010 ± 0.002CjGH5F XyG 0.040 ± 0.003 52.4 ± 0.8 74.8 ± 4.1Hydroxyethylcellulose (HEC) ND ND 0.090 ± 0.003Page 5 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 1,3/1,4-β-glucan, guar galactomannan, konjac glucoman-nan, beechwood xylan, wheat flour arabinoxylan, or xan-than for any of the three enzymes was observed. CjGH5D appeared to strictly require the branched XyG structure, while CjGH5E demonstrated trace activities against the artificial 1,4-β-glucans hydroxyethylcellulose (HEC) and carboxymethylcellulose (CMC) at the highest tested sub-strate concentration (2 mg mL−1); specific activities were 200 and 1500-fold less than XyG, respectively (Table 1). Similarly, CjGH5F was able to hydrolyze HEC with an 800-fold lower specific activity than XyG, while no activ-ity towards CMC was detected. Michaelis–Menten anal-ysis for XyG further underscored the high XyG specificity of the three enzymes: remarkably low Km values were observed and high kcat values recapitulated those previ-ously observed for predominant endo-xyloglucanases, including CjGH74 [22, 27, 36, 37] (Table  1, Additional file 1: Figure S6).Time-course analyses of native XyG polysaccharide hydrolysis products by HPAEC-PAD revealed that all three GH5_4 enzymes generated products of interme-diate retention time in the early stages of the reactions, with no significant generation of the  Glc4-based XXXG, XLXG, XXLG, and XLLG limit-digest products (Addi-tional file 1: Figures S7, S8; cf. Fig. 1). These results indi-cate that the three enzymes hydrolyze XyG through a dissociative, rather than processive [38] mechanism, and are thus canonical endo-xyloglucanases (EC; cf. EC, EC The limit-digest products fur-ther revealed that all C. japonicus GH5_4 enzymes spe-cifically catalyze hydrolysis at the anomeric position of the unbranched glucose residues of the (galacto)XyG pol-ysaccharide chain (Fig. 1). This cleavage pattern is typical for many GH5 [22, 34, 37], GH9 [39, 40], GH12 [34, 41–44], GH16 [45] and GH74 [46–50] endo-xyloglucanases, although certain GH5 [35, 36], GH7 [51], GH44 [40, 52] and GH74 [53–55] members preferentially hydrolyze the XyG backbone between branched glucosyl residues. The canonical XXXG-type XyGOs produced by CjGH5D, CjGH5E, and CjGH5F are direct substrates for the exo-glycosidases of the XyG gene cluster [13].With knowledge of the cleavage specificity of the GH5_4 members, we determined kinetic parameters for the hydrolysis of a panel of chromogenic oligosac-charides to reveal the contribution of side chain substi-tution on substrate recognition and catalysis (Table  2). All three enzymes were only weakly active on 2-chloro-4-nitrophenyl cellotrioside (GGG-β-CNP) and 2-chloro-4-nitrophenyl cellotetraoside (GGGG-β-CNP), with meagre increases in kcat/Km values arising from the addi-tion of potential − 4 subsite binding for the cellotetrao-side (Table  2, Additional file  1: Figure S9) (GH subsite nomenclature according to [56]). Strikingly, the addition of three α(1→6)-xylopyranosyl residues to the glucan backbone resulted in significant increases in catalytic effi-ciency for all GH5_4 members, which was manifested as 65-, 700-, and 150-fold higher kcat/Km values for XXXG-β-CNP vis-à-vis GGGG-β-CNP with CjGH5D, CjGH5E, and CjGH5F, respectively (Table 2, Additional file 1: Fig-ure S9). These values correspond to 11, 18, and 13  kJ/mol, respectively, of additional transition state stabiliza-tion in the formation of the covalent glycosyl-enzyme in these anomeric-configuration-retaining GH5 enzymes (calculated using the formula: ΔΔG‡ = − RT ln[(kcat/Km XXXG)/(kcat/Km GGGG)]) [57]. With XLLG-β-CNP, the specificity constants (kcat/Km) were only increased by 1.5 to five  fold for the three endo-xyloglucanases, thereby indicating that extending β(1→2)-galactopyranosyl residues (Fig.  1) have little additional effect on catalysis (Table 2).Table 2 Kinetic parameters of CjGH5_4 enzymes for (xylo)gluco-oligosaccharide glycosidesND not determined due to limited availability of substrateEnzyme catalytic domains Substrate Km mM kcat min−1 kcat/Km min−1 mM−1CjGH5D GGG-CNP ND ND 2.21 ± 0.05GGGG-CNP ND ND 5.36 ± 0.07XXXG-CNP 0.81 ± 0.10 281 ± 12 347 ± 45XLLG-CNP 0.18 ± 0.02 162 ± 4 900 ± 103CjGH5E GGG-CNP 11.8 ± 0.6 191 ± 7 16.2 ± 1.0GGGG-CNP 5.02 ± 0.35 180 ± 7 35.9 ± 2.8XXXG-CNP 0.010 ± 0.001 254 ± 4 (25.4 ± 2.6) × 103XLLG-CNP 0.010 ± 0.001 332 ± 9 (33.2 ± 3.4) × 103CjGH5F GGG-CNP ND ND 6.45 ± 0.30GGGG-CNP ND ND 16.5 ± 1.0XXXG-CNP 0.07 ± 0.01 169 ± 4 (2.41 ± 0.35) × 103XLLG-CNP 0.030 ± 0.002 393 ± 8 (13.1 ± 0.9) × 103Page 6 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 Covalent labelling of CjGH5D with an active‑site‑directed inhibitorActivesite affinity-based inhibitors are important tools for the detailed kinetic analysis of GH enzymes [58]. In particular, N-bromoacetylglycosylamine derivatives of xyloglucan oligosaccharides have been previously dem-onstrated to be specific active-site affinity labels for endo-xyloglucanases [36, 59]. A time- and concentra-tion-dependent inactivation of the enzyme CjGH5D was observed upon incubation with XXXG-NHCOCH2Br, which followed pseudo-first-order kinetics (Fig.  2). The dissociation constant Ki and the irreversible inactivation constant ki for XXXG-NHCOCH2Br  towards CjGH5D were 1.78 ± 0.17 mM and 0.17 ± 0.01 min−1, respectively, resulting in a ki/Ki value (9.3 × 10−2 mM−1 min−1) that was comparable to that previously observed for a Prevo-tella bryantii GH5_4 member (PbGH5A) [36]. Notably, intact protein mass spectrometry of CjGH5D following incubation with the inhibitor indicated covalent label-ling with 1:1 stoichiometry and no over-labelling of the enzyme (Additional file 1: Figure S10).CjGH5D crystallographyA tertiary structure of the catalytic domain of CjGH5D was determined at 1.6 Å resolution in uncomplexed “apo” form by X-ray crystallography and molecular replace-ment with BoGH5A (pdb: 3ZMR, [22]). The overall struc-ture of CjGH5D (residues Gly96 to Gln468) is an (β/α)8 barrel as is typical for GH5 family members (Fig.  3a). Despite sequence identities in the 25–40% range, the structure is similar to the catalytic domains of many GH5 enzymes, most of which are annotated as xyloglucanases, glucanases and lichenases, with typical alignment values of approximately 310 residues aligning with an rmsd of 1.3 Å [60]. For example, the structure used for molecu-lar replacement, BoGH5A, overlaps with an r.m.s.d. of 1.1  Å over 332 equivalent Cα atoms with 40% identity. Minor differences are observed between the two struc-tures in loops at the end of core helices. BoGH5A has an extra loop Val170–Gly180 (residues equivalent to Ile137–Gly138 in CjGH5D) which enables the formation of a hydrogen bond to the − 4′-xylosyl residue of ligand XXXG (between N Val182 and the sugar ring O atom, vide infra).A 1.9  Å-resolution product complex of CjGH5D was obtained by soaking crystals with a mixture of  Glc12-based XyGOs of variable sidechain galactosylation. Time (min)[I] (mM)v o(M min–1)k app(min–1)0 20 40 60 80 1000 2 4 6 8 10 12 14 16 1802550751000. mM0.5 mM1.0 mM2.0 mM4.0 mM8.0 mM12 mM16 mMABFig. 2 Inhibition kinetics of CjGH5D with XXXG-NHCOCH2Br. a Initial-rate enzyme activity over time (single determinations). b Pseudo-first-order rate constants (kapp) obtained from the fitted curves shown in a. Bars represent errors in kapp values from curve-fitting. The 95% confidence interval is indicated (pink band) for the fitted curve (solid line)Fig. 3 Three-dimensional structure of CjGH5D in complex with XXXG-NHCOCH2Br and XyGOs. a Cartoon representation of the secondary structure of CjGH5D colour ramped from the N-terminus (blue) to the C-terminus (red). The two ligands XXXG-NHCOCH2Br and GXLG are overlaid in the active site cleft and shown in green and magenta sticks, respectively. b A close-up view of the active site cleft with the overlaid ligands XXXG-NHCOCH2Br in green and XXLG in magenta showing different amino acids interacting with the carbohydrate ligands. c 2Fo − Fc (σA/maximum likelihood weighted) electron density contoured in blue around GXLG in the CjGH5D-XXLG complex (left panel) and the chemical structure of the corresponding ligand (Right panel). Insufficient electron density was observed for the − 4′ xylosyl residue to allow modelling, therefore, it is shown in grey. d 2Fo − Fc electron density at 1σ (approx. 0.2 e−/Å3) contoured in blue around XXXG-NHCOCH2 moiety in the CjGH5D-XXXG-NHCOCH2Br complex (left panel) and chemical structure of the corresponding ligand (right panel). The bromide leaving group is shown in grey(See figure on next page.)Page 7 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 Here, we anticipated that the substrate mixture would be hydrolyzed and that the enzyme would selectively bind the oligosaccharide for which it had the best affinity. Commensurate with limit-digest analysis, we observed a  Glc4-based oligosaccharide backbone spanning the −  4 to −  1 subsites for both molecules in the asymmetric unit: GXLG in molecule A (with glucose in the − 4 sub-site and the − 3′-xylosyl group modelled at occupancies acdbO OHOOHOOHOOHO OHOOHOHOOHOOHOOHOOHOHHOOHOHOOOHOOHOHHOHN BrOO OHOOHOOHOOHO OHOOHOHOOHOOHOHOHHOOOH OOHHOOHOHOOOHOOHOHOHOOHHOOPage 8 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 of 0.5 and 0.7, respectively) and GXXG in molecule B (here, there was insufficient electron density in the Fo − Fc difference map to allow unambiguous modelling of a galactose on the − 2′-xylosyl unit). In the − 1 sub-site, the glucosyl residue interacts with the catalytic acid base Glu255 (via O1), and nucleophile Glu390 via O2. In addition, O3 is hydrogen bonded to His208. In molecule B, the equivalent glucose also hydrogen bonds via O2 to Asn254 and His208 (Fig. 3b, c).A second oligosaccharide complex was obtained at 2.1  Å resolution by soaking CjGH5D crystals with the N-bromoacetyl affinity label XXXG-NHCOCH2Br, in which the reagent had indeed reacted through attack of the catalytic general acid/base sidechain to displace the bromide nucleofuge. In molecule A of the asymmetric unit, there is electron density for GXXG-NHCOCH2-CjGH5D, whilst in molecule B, XXXG-NHCOCH2-CjGH5D is modeled, but with the − 3′- and − 4′-xylosyl sugars modelled at half occupancy. The carboxyl oxy-gen of the N-acetyl moiety forms a hydrogen bond with His323. There are hydrogen bonds between this sub-site −  1 sugar and the catalytic nucleophile Glu390, and also to His208 and Asn254 (Fig.  3b, d). These are similar to interactions observed in the structure of an analogous XXXG-NHCOCH2-PbGH5A complex struc-ture (pdb: 5D9P, [36]). Glucose in the −  2 subsite is hydrogen bonded via O3 to ND2 Asn132 and via O2 to NE1 Trp432; this latter interaction is notably long (ca. 3.2  Å), which may reflect the positioning of the trypto-phan as the − 1 subsite stacking residue. The equivalent Asn/Trp interactions are also seen in related enzymes: Asn28 and Trp324 in the XXXG-NHCOCH2-PbGH5A complex (pdb: 5D9P) and Asn165 and Trp472 in the BoGH5A-XXXG complex (PDB 3ZMR). In addition to Trp432, Trp143 provides aromatic stacking interac-tions with the glucose in the − 3 subsite (homologous to Trp324 and Trp48 in PbGH5A, and Trp472 and Trp185 in BoGH5A, respectively), while Trp209 lies against the −  2′-xylosyl residue (as does the equivalent Trp252 in BoGH5A). This pattern of conserved/highly invariant residues interacting with the xyloglucan chain presum-ably accounts for the fact that despite sharing amino acid identity as low as 30%, these enzymes are all tailored for xyloglucan as a substrate (Fig.  4). None of these three GH5D structures exhibits direct interactions with glu-cosyl units in the − 3 and − 4 subsites with the protein. The − 3′-xylosyl unit is tethered by two hydrogen bonds between O3 and O4 and Asp438, which hold the sugar perpendicular to the orientation of the equivalent xylose in the XXXG-NHCOCH2-PbGH5A and BoGH5A:XXXG complexes (in the latter, the xylose lies parallel to the side chain of Tyr476).The covalent adduct formation through the reactivity of the N-bromoacetyl reagent is fascinating given that in the structures observed here, the attack is made by the acid–base Glu255, as opposed to the enzymatic nucleophile of the enzymatic reaction, Glu390. This latter residue is poised for nucleophilic attack at the anomeric carbon, C1, of the − 1 subsite glucoside. However, Glu390 is too distant (6–7 Å) from the reactive carbon of the N-bro-moacetyl moiety, and has impossible geometry and steric hindrance, to permit nucleophilic interception. The reac-tive group, however, is located in the + 1 subsite—some 3.8 Å from C1—thus can be fortuitously attacked by the acid/base which is in almost ideal position for SN2 attack on the reactive carbon to displace the bromide. Such Fig. 4 Divergent (wall-eyed) stereo surface representation of CjGH5D-GXLG showing regions of sequence conservation. Surfaces of conserved and non-conserved residues, shown in purple at reduced opacity and sea-green, respectively, were calculated from an amino acid sequence alignment of GH5 domains of CjGH5D, CjGH5E, CjGH5F and five additional GH5 members showing E.C. activity (Additional file 1: Figure S4). Figure was generated using CCP4MG [81]Page 9 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 a reaction is facilitated, either prior, or subsequent to attack by rotation around the CB–CG bond, which leaves the side-chain in a different rotamer after the reaction relative to its “normal” position in unreacted complexes (Fig. 3d).Mutational analyses of C. japonicus GH5_4 genes indicate a complex mode of action for the initial stages of xyloglucan degradationFacilitated by knowledge of their broadly similar catalytic properties, we embarked on a comprehensive reverse-genetic analysis in an attempt to delineate the biological functions of the individual GH5_4 and GH74 endo-xylo-glucanases, using recently developed in-frame gene dele-tion techniques [9].In-frame deletion mutants were first generated in the XyG gene cluster encoding the three exo-glycosidases and the TBDT (Fig.  1b) to provide benchmark con-trols for subsequent analysis of endo-xyloglucanase deletion mutants. Recapitulating our previous work using insertional mutants [13], an in-frame Δxyl31A (α-xylosidase) mutant was unable to grow on XyG due to an inability to remove non-reducing terminal xylosyl residues as the first essential step in XyGO saccharifi-cation (Fig.  5a, cf. Fig.  1). A ΔCJA_2709 (TBDT) single mutant strain had a significant growth defect, presum-ably resulting from a decreased ability to uptake extracel-lularly produced XyGOs into the periplasm. The deletion of bgl35A also attenuated growth, due to an inability of the strain to access the full complement of sidechain monosaccharides. As expected, growth of the Δafc95A (α-l-fucosidase) mutant on tamarind (galacto)xyloglucan was identical to the wild-type strain, because this read-ily available substrate lacks the terminal fucosyl residues typically found in dicot primary cell wall XyG (Fig.  1a). Moreover, all XyG gene cluster mutant strains grew simi-larly to wild type in glucose containing medium (Addi-tional file 1: Figure S11).With these control experiments complete, we next ana-lyzed the effect of deleting the individual GH5_4- and 0 5 10 15 20 - GH5/GH74 triple/quad mutantsTime [hours]OD600Wild Type3010 3337 29593010 3337 2959 2477XyG - GH5/GH74 double mutants0 5 10 15 20 [hours]OD600Wild Type3010 33373010 29593010 24773337 29592959 24773337 2477XyG - GH5/GH74 single mutants0 5 10 15 20 [hours]OD600 Wild Type3010333729592477XyG - XyGUL0 5 10 15 20 [hours]OD600Wild Typeafc95Abgl35Axyl31A2079A BC DFig. 5 Growth analysis of in-frame deletions of GH5_4, and GH74 mutant strains on xyloglucan. a Control experiment with XyGUL in-frame deletion mutant strains. b Single, c double, d triple and quadruple deletion mutants were made with the GH5_4 and GH74 genes; CJA_3010 encodes CjGH5D, CJA_3337 encodes CjGH5E, CJA_2959 encodes CjGH5F, and CJA_2477 encodes CjGH74. Graphs represent the average of three biological replicates and error bars represent the standard deviation. All strains grew similarly to wild-type when grown with MOPS-glucose defined medium (Additional file 1: Figure S12)Page 10 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 GH74-encoding genes. Despite original indications by RNAseq analysis of a potential lead role for CjGH5D in XyG utilization, in-frame deletion of CJA_3010 surpris-ingly did not elicit a statistically significant growth defect (Fig.  5b). Likewise, strains containing single in-frame deletions of CJA_3337, CJA_2959, and CJA_2477 grew identically to the wild-type strain. Moreover, compre-hensive combinatorial mutagenesis did not yield a strain with a substantial growth defect for any combination of double, triple, or quadruple mutants (Fig.  5c, d). Fur-thermore, the growth traits (maximum OD and growth rate) of both the wild-type and the quadruple deletion mutant were similar when reduced (0.25%) or limiting (0.125%) concentrations of XyG were used (Additional file 1: Figure S12 and Table S6). Collectively, these results suggest that despite their individual high activities and specificities toward XyG, as defined by the biochemical and structural analyses described above, the GH5_4 and GH74 members are not the only enzymes encoded by C. japonicus with sufficient xyloglucanase activity to sup-port growth.CjGH5D is predicted to be attached to the exterior face of the outer membrane by N-terminal lipidation, while CjGH5E, CjGH5F, and CjGH74 are likely to be secreted enzymes (vide supra). Thus, we hypothesized that other secreted enzymes, with either predominant or side hydrolytic activities toward XyG, may be enabling growth of the quadruple mutant. Deletion of the type-two secre-tion system (T2SS) in the Δgsp mutant has been previ-ously shown to abolish the ability of C. japonicus to secrete cellulases [11], and constitutes a powerful tool to restrict extracellular secretion of CAZymes in general. Interestingly, the introduction of the CJA_3010 deletion into the Δgsp background resulted in only limited growth attenuation on xyloglucan (Additional file 1: Figure S13). With the T2SS extracellular secretion pathway disabled, the ability of the Δgsp ΔCJA_3010 strain to grow on XyG strongly suggests the presence of other membrane-bound XGases that effect XyG depolymerization in a physiologi-cally relevant manner. These data sharply contrast obser-vations for the human gut symbiont Bacteroides ovatus, for which deletion of a single GH5_4 member from the XyGUL resulted in complete loss of growth on XyG [22].Predominant xyloglucanase activity has been demon-strated previously in members of CAZyme families GH5, GH7, GH9, GH12, GH16, GH44, and GH74, and poten-tially may constitute a side activity in other endo-β(1,4)glucanases (cellulases) [23, 61]. Examination of  the C. japonicus genome indicates the presence of multiple GH5 (n = 15), GH9 (n = 3), and GH16 (n = 9) encod-ing genes, in addition to the single GH74 member ([8]; for a summary table, see http://www.cazy.org/b776.html). Further, Deboy et  al. [8] predicted that there are approximately 45 membrane-bound CAZymes. Although it constitutes a significant undertaking that is beyond the scope of the present study, our future investigations will focus on scrutinizing these additional CAZymes in the context of XyG utilization by C. japonicus.ConclusionsWe previously proposed a model of XyG utilization by C. japonicus, in which an extracellular endo-xyloglucanase mediates degradation of the polysaccharide to XyGOs for uptake via the TBDT, followed by complete hydrolysis to monosaccharides in the periplasm by the exo-glycosi-dases encoded by the XyG gene cluster [13]. Our pre-sent study, combining biochemical and reverse-genetic analyses, reveals that the number of actors in the initial cleavage event is significantly greater than what was orig-inally anticipated by bioinformatics. We propose that the existence of so many extracellular endo-xyloglucanases of apparently overlapping biochemical function can be explained by a physiological interplay of secreted recon-naissance enzymes and cell-surface-bound, proximal XyG degraders (Fig. 1b).Thus, the secreted GH74 and two secreted GH5_4 enzymes may act as highly mobile, primary “unravellers” of the plant cell, liberating XyG fragments from the lig-nocellulose matrix (Fig. 1c). Indeed, the concept of “sens-ing” polysaccharidases playing a lead role in generating inducers has been previously proposed [62]. As plant cell wall polysaccharide degradation advances, more inti-mate contact between the bacterial cell surface and the substrate may ensue, engaging the outer-membrane-bound CjGH5D and a more efficient interplay between XyG backbone hydrolysis and direct TBDT-mediated uptake of the oligosaccharide products. The coordinated capture, hydrolysis, and uptake of partially hydrolyzed polysaccharides as a successful competitive strategy has considerable precedent in the Polysaccharide Utilization Loci of the Bacteroidetes [63]. Moreover, the need to ini-tiate cell wall “unravelling” has been suggested to explain why saprophytes such as C. thermocellum, which are unable to utilize xyloglucan or xylan for growth, contain endo-xyloglucanases and endo-xylanases within their cel-lulosomes [39, 64].MethodsTranscriptomic analysisRNAseq sampling and analysis was performed as previ-ously described [9, 12]. Briefly, C. japonicus cultures were grown in 500  mL flasks at 30  °C shaking at 200 RPM.  OD600 was measured every hour to monitor growth and samples were taken during exponential and station-ary phase. Within 2  min of sampling, metabolism was stopped using a phenol/ethanol solution (5%/95%). The Page 11 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 samples were immediately pelleted by centrifugation at 8000 g at 4 °C for 5 min. The supernatant was discarded and cell pellets were then flash frozen using a dry ice/ethanol bath and stored at −  80  °C. RNA extraction, library preparation, and sequencing was performed by GeneWIZ (South Plainfield, NJ). Illumina HiSeq  2500 was performed as 50  bp single-reads with at least 10 million reads generated per sample. The raw data have been submitted to the NCBI Gene Expression Omnibus (Accession GSE109594).Bioinformatic analysisThe full-length proteins encoded by ORFs CJA_3010 (CjGH5D), CJA_3337 (CjCBM2-CBM10-GH5E) and CJA_2959 (CjFN3-GH5F) in C. japonicus genome were screened for the presence of a signal peptide using Sig-nalP 4.0 [32] and LipoP 1.0 [65]. The modular architec-ture of the three enzymes was obtained from BLASTP analysis and additional alignment with representative GH and CBM modules from the CAZy Database [7] using ClustalW [66].Cloning of cDNA encoding protein modulescDNA encoding the full-length enzymes CjSRL-GH5D, CjCBM2-CBM10-GH5E and CjFN3-GH5F, in addition to the catalytic domains CjGH5D, CjGH5E and CjGH5F were PCR amplified from C. japonicus genomic DNA; all constructs were designed such that the native predicted signal peptide was removed (PCR primers are listed in Additional file 1: Table S7). The amplified CjSRL-GH5D, CjCBM2-CBM10-GH5E, CjFN3-GH5F, CjGH5D and CjGH5F products were double-digested with NheI and XhoI, gel purified and ligated to the respective sites of pET28a to fuse an N-terminal 6× His-Tag. The ampli-fied CjGH5E product was ligated in an SspI linearized pMCSG53 vector using the Ligation Independent Clon-ing (LIC) strategy [67]. Successful cloning was confirmed by PCR and plasmid DNA sequencing. Q5 high fidelity DNA polymerase was used for all the PCR amplifications.Gene expression and protein purificationConstructs were individually transformed into the chemically competent E. coli Rosetta DE3 cells. Colonies were grown on LB solid media containing kanamycin (50 µg mL−1) and chloramphenicol (30 µg mL−1) [CjSRL-GH5D, CjCBM2-CBM10-GH5E, CjFN3-GH5F, CjGH5D and CjGH5F], or containing ampicillin (50 µg mL−1) and chloramphenicol (30 µg mL−1) [CjGH5E]. One colony of the transformed E. coli cells was inoculated in 5  mL of LB medium containing the same antibiotics and grown overnight at 37  °C (200  rpm). The whole overnight cul-ture was used to inoculate 500 mL of TB liquid medium containing the appropriate antibiotics. Cultures were grown at 37  °C (200 rpm) until D600 = 0.6. Overexpres-sion was induced by adding IPTG to a final concentra-tion of 0.1  mM. After induction, cultures were grown overnight at 16  °C (200  rpm). Cultures were then cen-trifuged (4220 g at 4 °C) and pellets were resuspended in 10  mL of E. coli lysis buffer containing 20  mM HEPES, pH 7.0, 500  mM NaCl, 40  mM imidazole, 5% glycerol, 1 mM DTT and 1 mM PMSF. Cells were then disrupted by sonication and the clear supernatant was separated by centrifugation at 4 °C (24,000 g for 45 min). Recombi-nant proteins were purified from the clear soluble lysates using a  Ni+2-affinity column utilizing a gradient elution up to 100% elution buffer containing 20 mM HEPES, pH 7.0, 100 mM NaCl, 500 mM imidazole, and 5% glycerol in an FPLC system. Purity of the recombinant proteins was determined by visualizing the protein contents of the fractions on SDS-PAGE. Pure fractions were pooled, concentrated, and buffer exchanged against 50  mM phosphate buffer (pH 7.0) containing 10% glycerol. Pro-tein concentrations were then determined using Epoch Micro-Volume Spectrophotometer System  (BioTek®, USA) at 280  nm, and identities of the purified proteins were confirmed by intact mass spectrometry [68]. Puri-fied proteins were then aliquoted and stored at − 80  °C until needed.Carbohydrate sourcesTamarind seed XyG, konjac glucomannan (KGM), barley β-glucan (BBG), wheat flour arabinoxylan, and beech-wood xylan were purchased from  Megazyme® (Bray, Ireland). Hydroxyethylcellulose (HEC) was purchased from  Amresco® (Solon, USA). Carboxymethyl cellulose was purchased from Acros Organics (New Jersey, USA). Guar gum was purchased from Sigma  Aldrich® (St. Lou-ise, USA). Xanthan gum was purchased from  Spectrum® (New Brunswick, USA). 2-Chloro-4-nitrophenyl (CNP)-β-d-cellotrioside (GGG-β-CNP) and CNP-β-d-cellotetraoside (GGGG-β-CNP) were purchased from  Megazyme®. XXXG-β-CNP and XLLG-β-CNP were pre-pared as previously described [69].  Glc4-based XyGOs (XXXG, XLXG, XXLG, and XLLG; nomenclature accord-ing to [20]) and  Glc8-based XyGOs were prepared from XyG powder (Innovassynth Technologies, Maharashtra, India) as previously described [61]. XXXG-NHCOCH2Br was synthesized as previously described [59].Carbohydrate analyticsHigh Performance Anion-Exchange Chromatography with Pulsed Amperometric Detection (HPAEC-PAD) was performed on a Dionex ICS-5000 DC HPLC system operated by the Chromeleon software version 7 (Dionex) using a Dionex Carbopac PA200 column. Solvent A was double-distilled water, solvent B was 1  M sodium Page 12 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 hydroxide (NaOH), and solvent C was 1 M sodium ace-tate (NaOAc). The gradient used was: 0–4 min, 10% sol-vent B and 2.5% solvent C; 4–24 min, 10% B and a linear gradient from 2.5 to 25% C; 24–24.1 min, 50% B and 50% C; 24.1–25  min, an exponential gradient of NaOH and NaOAc back to initial conditions; and 25–31 min, initial conditions.Matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) was performed on a Bruker Dal-tonics Autoflex System (Billerica, USA). The matrix, 2,5-dihydroxy benzoic acid, was dissolved in 50% meth-anol in water to a final concentration of 10  mg  mL−1. Oligosaccharide samples were mixed 1:1 (v/v) with the matrix solution. One microliter of this solution was placed on a Bruker MTP 384 ground steel MALDI plate and left to air dry for 2 h prior to analysis.Enzyme kinetic analysisAll enzyme activities toward polysaccharides were deter-mined using a bicinchoninic acid (BCA) reducing-sugar assay [70]. The effect of temperature on xyloglucanase activity was determined by incubating the recombinant catalytic domain: CjGH5D (0.098 µg), CjGH5E (0.086 µg), CjGH5F (0.017  µg) with tamarind seed xyloglucan at a final concentration of 1  mg  mL−1. Citrate buffer (pH 6, CjGH5D and CjGH5F) or phosphate buffer (pH 7.5, CjGH5E) was used to a final concentration of 50 mM in a total reaction volume of 200 µL. Reaction mixtures were incubated for 10  min at temperatures ranging from 25 to 80  °C prior to the BCA assay. To determine the pH-rate profile, the same XyG concentration was incubated with the same enzyme amounts, except for CjGH5D (0.049  µg), for 10  min at 50  °C (CjGH5D, CjGH5F), or 55  °C (CjGH5E), with 50 mM final concentration of the following buffers: citrate (pH 3–6.5), phosphate (pH 6.5–8), and glycine (pH 8.5–9).For qualitative activity assessment against the other polysaccharide substrates, 1  µg of each recombinant enzyme was added to XyG, HEC, CMC, BBG, KGM, wheat flour arabinoxylan, beechwood xylan, xanthan gum, and guar gum to a final concentration of 2 mg mL−1 in 200 µL reaction volumes containing 50 mM phosphate buffer (pH 7.5: CjGH5D and CjGH5E, or pH 7: CjGH5F). Mixtures were then incubated at 50  °C (CjGH5D and CjGH5F) or 55 °C (CjGH5E) for 10 min before the gener-ated reducing ends were detected using BCA assay.To determine specific activity values of CjGH5 enzymes toward XyG, final concentration of 0.75, 2.59, and 0.71 nM of the recombinant purified catalytic mod-ules CjGH5D, CjGH5E, and CjGH5F, respectively, was incubated with tamarind seed XyG (1  mg  mL−1) in 200 µL reaction mixtures containing 50  mM phos-phate buffer (pH 7.5: CjGH5D and CjGH5E, or pH 7: CjGH5F). Likewise, specific activity values of CjGH5 enzymes toward HEC were obtained by incubating CjGH5E and CjGH5F at a final concentration of 1.04 and 0.65  µM, respectively, with 2  mg  mL−1 HEC in 200  µL reaction mixtures containing 50  mM phosphate buffer (pH 7.5: CjGH5E or pH 7: CjGH5F). For specific activ-ity toward CMC, final concentration of 1.04  µM of the purified catalytic module CjGH5E was incubated with CMC (2  mg  mL−1) in 200  µL reaction volume contain-ing 50 mM phosphate buffer (pH 7.5). All reaction mix-tures were incubated at 50 °C (CjGH5D and CjGH5F) or 55 °C (CjGH5E) for 10 min prior to the BCA assay and all assays were performed in triplicates.To determine Michaelis–Menten parameters for XyG, eight different concentrations of XyG solutions were used over the range 0.025–1  mg  mL−1. The recombi-nant enzymes CjGH5D (0.007  µg), CjGH5E (0.022  µg), and CjGH5F (0.006 µg) were individually incubated with each XyG concentration at 50 °C (CjGH5D and CjGH5F) or 55  °C (CjGH5E) for 10  min in a 200  µL final reac-tion mixture containing 50  mM phosphate buffer (pH 7.5: CjGH5D and CjGH5E, or pH 7: CjGH5F). Km and kcat values were determined by non-linear fitting of the Michaelis–Menten equation to the data in  Sigmaplot® (Systat software Inc.)To identify Michaelis–Menten constants for the chro-mogenic substrates, different dilution series were estab-lished to give final concentration ranges of 0.0625–8 mM (GGG-β-CNP), 0.0625–8  mM (GGGG-β-CNP), 0.002– 4 mM (XXXG-β-CNP), and 0.002–2 mM (XLLG-β-CNP). Substrate mixtures (225  µL) containing 50  mM phos-phate buffer with the optimum pH of the enzyme were pre-incubated for 10  min at the optimum temperature of the enzyme (vide supra). 25  µL of 10× CjGH5D (to give 30–3800  nM final concentration according to the tested substrate), CjGH5E (3–260  nM), and CjGH5F (4- 650 nM) was added to the substrate mixtures before the release of the aglycone was continuously monitored by measuring the change in absorbance at 405  nM for 2  min in a Cary50 UV–visible spectrophotometer (Var-ian). CNP molar extinction coefficients were determined to be 17,288 M−1 cm−1 in 50 mM phosphate buffer pH 7 and 17,741 M−1 cm−1 in 50 mM phosphate buffer pH 7.5.Enzyme product analysisTo determine the limit-digest products of the CjGH5 enzymes, 5 µg of each recombinant enzyme was incubated with tamarind seed XyG at final concentra-tion of 0.25  mg  mL−1 for 7  h (40  °C) in a 200  µL reac-tion mixture that contained 50 mM phosphate buffer of the optimum pH of the tested enzyme (pH 7.5: CjGH5D and CjGH5E, or pH 7: CjGH5F). The reaction mix-ture was then diluted 5 times prior to product analysis Page 13 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 by HPAEC-PAD. To determine the mode of action of the enzyme, 0.01  µg of CjGH5  enzymes was incubated at 40  °C with 1  mg  mL−1 final concentration of tama-rind seed XyG in 200  µL reaction volumes containing the same buffers used in the  limit-digest analysis. The reaction was stopped at different time points by adding 100  µL of 1 M  NH4OH. Reaction mixtures were then diluted 2 times with water prior to product analysis by HPAEC-PAD.Inhibition kinetics and active‑site labellingInhibition kinetic parameters were determined as pre-viously described [59]. Briefly, a final concentration of 0.23 μM of CjGH5D was incubated with a series of differ-ent concentrations (0.5–16 mM) of XXXG-NHCOCH2Br at 40  °C in 20  mM phosphate buffer (pH 7.5) for up to 90 min. BSA to a final concentration of 0.1 mg mL−1 was added to the inhibition mix to prevent the non-specific loss of activity. Small samples (10  µL) of the incubate were periodically diluted 1:100 in 20  mM phosphate buffer (pH 7.5), and 100  µL of the diluted incubate was added to 100  µL of the pre-incubated substrate XXXG-CNP at 40  °C (0.1  mM final substrate concentration in the assay). Residual activities of the enzyme were determined by measuring the rate of the release of the chromophore 2-chloro-4-nitrophenolate [69] at 405  nm in Agilent Cary 60 UV–vis Spectrophotometer. Initial-rate kinetics were measured in the strictly linear range of the enzyme. Equations 1 and 2 were used to determine Ki and ki values by non-linear regression curve-fitting using OriginPro 2015 software as previously described [59].Intact protein masses were determined on a Waters Xevo Q-TOF with a nanoACQUITY UPLC system, according to the method previously published [68], with 2.5 mM inhibitor and 4.52 μM of enzyme.Construction of C. japonicus mutants and growth conditionsIn-frame deletion mutants were made as previously described [9]. Briefly, 500  bp regions up- and down-stream of the genes of interest were amplified by PCR (CJA_3010, CJA_3337, and CJA_2959) or synthesized by GeneWIZ (South Plainfield, NJ) (CJA_2477) and assembled into pK18mobsacB by the method of Gibson et al. [71]. Deletions were confirmed by PCR. For a com-plete list of primers used see Additional file 1: Table S8. Cultures were grown at 30  °C with 200 RPM shaking in (1)V = V0exp(−kappt)+ yoffset(2)kapp =ki[I]Ki + [I]MOPS minimal media containing 0.25% (w:v) glucose, 0.25%   (w:v) XyG  or 0.5% (w:v)  XyG as the sole carbon source in 18 mm test tubes or in 96 well flat bottom poly-styrene plates (Corning). Growth was measured using a Spec20D+ spectrophotometer (Thermo Scientific) or a Tecan Plate reader (Tecan, Switzerland). All experi-ments were performed in biological triplicate. Statistical analysis was performed using Graphpad Prism 6 software package (La Jolla, CA) where appropriate.Crystallization, X‑ray crystallography and structure solutionCjGH5D was crystallized in sitting drops by the Vapour Diffusion method, using protein at 21 mg mL−1 in 50 mM sodium citrate pH 6.5 and 10% glycerol over a well solu-tion comprised of 1.9  M ammonium sulfate, 0.1  M 2-(N-morpholino)ethanesulfonic acid (MES) pH 5.5. The drop consisted of 0.5 μL enzyme, 0.1 μL seed stock and 0.4 μL well solution, and the seed stock was prepared by vortexing crystals, grown over 1.4 M ammonium sulfate, 0.1 M MES pH 5.5, 1% (w/v) polyethylene glycol 1000, in an Eppendorf tube with a polystyrene bead. Crystals were harvested into liquid nitrogen using nylon CryoLoops™ (Hampton Research). A non-ligand complexed “apo” dataset was collected from a crystal after immersing for a few minutes in a cryoprotectant solution comprised of the mother liquor supplemented with 20% (v/v) glycerol. Data were collected at Diamond beamline I04, and pro-cessed using DIALS [72], and scaled using AIMLESS [73] to 1.6 Å. The space group is  P212121 with unit cell dimen-sions 55.0, 96.4, 159.0 Å, and there are 2 molecules in the asymmetric unit.The structure was solved by molecular replacement using Phaser [74] using residues 138–500 of PDB entry 3zmr as the search model [22], which align to residues 106–464 of CjGH5D, with which they share 38% identity (using the program lalign from the FASTA package [75]). The structure was built automatically using Buccaneer [76] and refined using cycles of manual model rebuilding using Coot [77] followed by refinement with REFMAC [78], including cycles using anisotropic B-factor refine-ment. In addition to the two protein chains, there are 20 molecules of glycerol, 5 sulfate ions and 3 molecules of PEG (introduced from the seed stock solution). Data collection and refinement statistics for all structures are given in Additional file 1: Table S9.A crystal of CjGH5D, grown as above over a well solution comprised of 2.3 M ammonium sulfate, 0.1 M MES pH 5.5, was soaked for 27.5  h in 1.85  M ammo-nium sulfate, 0.1 M MES pH 5.5 with 4.5 mM XXXG-NHCOCH2Br, and fished directly into liquid nitrogen. Data were collected at Diamond beamline I03, and pro-cessed using DIALS [72]. After scaling with AIMLESS Page 14 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 [73], the data were cut off at a resolution of 2.1  Å, as although the X-ray images showed significant spot smearing, the Rmerge and  CC1/2 values were good (6.2% overall, 54.7% in the outer shell and 0.998 overall, 0.933 in the outer shell, respectively).Crystals grown under similar conditions, over a well containing 1.6  M ammonium sulfate, 0.1  M MES pH 5.5, were soaked in the presence of a mixture of  Glc12-based XyGOs (produced as described in [46] at a concentration of 5 mM for 5 h, before fishing into liq-uid nitrogen via a cryoprotectant solution, as for the apo crystal. A dataset was collected at Diamond beam-line I04 and processed using DIALS and scaled using AIMLESS to 1.9 Å.Both ligand structures were solved initially using the apo structure as a model for REFMAC, and the ligand was placed after the protein chain had been rebuilt (using cycles of Coot interspersed with refinement in REFMAC) and some water molecules added. All models were vali-dated using MolProbity [79] and the sugar conformations of the ligand in the complex structures were checked using Privateer [80]. Problems with diffraction anisotropy in both ligand datasets limited the possibility of refin-ing the structures to R/Rfree lower than 0.22/0.28 and 0.23/0.30 for the complexes with XXXG-NHCOCH2 and GXLG (produced after the  Glc12 soak), respectively.AbbreviationsXyG: xyloglucan; XyGO: xylogluco-oligosaccharides; CAZy: carbohydrate active enzymes; GH5: glycoside hydrolase family 5; CBM: carbohydrate binding module; SRL: serine rich linker; HPAEC-PAD: high performance anion exchange chromatography-pulsed amperometric detector; MALDI-TOF: matrix assisted laser desorption ionization-time of flight; KGM: konjac glucomannan; BBG: barley β-glucan; HEC: hydroxyethylcellulose; CNP: 2-chloro-4-nitrophenyl; BCA: bicinchoninic acid; TBDT: TonB-dependent transporter; XyGUL: xyloglucan utilization locus; MES: 0.1 M 2-(N-morpholino)ethanesulfonic acid.Authors’ contributionsMA performed the bioinformatic analysis, recombinant protein production, and biochemical characterization. CN performed the RNAseq analysis and all mutational analysis. NJ synthesized the active-site affinity label and, together with MA, performed inhibition and protein MS analysis. JGG designed the RNAseq and mutational analysis experiments, and assisted in analyzing the RNAseq and mutational data. WAO solved X-ray structures which were ana-lysed under supervision of GJD. HB devised the overall study and supervised research. All authors contributed to writing the article. All authors read and approved the final manuscript.Author details1 Michael Smith Laboratories, University of British Columbia, 2185 East Mall, Vancouver, BC V6T 1Z4, Canada. 2 Department of Chemistry, University of Brit-ish Columbia, 2036 Main Mall, Vancouver, BC V6T 1Z1, Canada. 3 Department of Biological Sciences, University of Maryland, Baltimore County, Baltimore, MD 21250, USA. 4 Department of Chemistry, University of York, Heslington, Additional fileAdditional file 1. Supplemental Tables S1–S9 and Supplemental Figures S1–S13.York YO10 5DD, UK. 5 Department of Biochemistry and Molecular Biology, University of British Columbia, 2350 Health Sciences Mall, Vancouver, BC V6T 1Z3, Canada. 6 Department of Botany, University of British Columbia, 6270 University Blvd., Vancouver, BC V6T 1Z4, Canada. AcknowledgementsWe thank Diamond Light Source (Harwell, UK) for access to beamlines I03 and I04 (proposal mx9948) that contributed to the results presented here. Additionally, we thank E.C. Monge (Gardner Lab, UMBC) for assistance with C. japonicus growth experiments.Dedication: This article is dedicated to Cellvibrio japonicus vanguard Prof. Harry J. Gilbert on the occasion of his retirement.Competing interestsThe authors declare that they have no competing interests.Availability of data and materialsThe structural datasets generated during the current study are available in the RCSB protein repository under the following PDB IDs: CjGH5D: 5OYC, XXXG-NHCOCH2–CjGH5D: 5OYD, CjGH5D-GXLG: 5OYE (https ://www.rcsb.org/). The RNAseq datasets generated during the current study are available in the NCBI Gene Expression Omnibus (GEO, https ://www.ncbi.nlm.nih.gov/geo/) reposi-tory under Accession GSE109594. Other datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.Consent for publicationNot applicable.DisclaimerThis report was prepared as an account of work sponsored by an agency of the United States Government. Neither the United States Government nor any agency thereof, nor any of their employees, makes any warranty, expressed or implied, or assumes any legal liability or responsibility for the accuracy, completeness, or usefulness of any information, apparatus, product, or process disclosed, or represents that its use would not infringe privately owned rights. Reference herein to any specific commercial product, process, or service by trade name, trademark, manufacturer, or otherwise does not necessarily constitute or imply its endorsement, recommendation, or favor-ing by the United States Government or any agency thereof. The views and opinions of authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof.Ethics approval and consent to participateNot applicable.FundingWork at the University of British Columbia was generously supported by the Natural Sciences and Engineering Research Council of Canada (Discovery Grant and Strategic Network Grant for the “NSERC Industrial Biocatalysis Net-work”, http://www.ibnet .ca/), the Michael Smith Laboratories (faculty start-up and post-graduate funding), the Canada Foundation for Innovation, and the British Columbia Knowledge Development Fund. Work at UMBC was sup-ported by the US Department of Energy, Office of Science, Office of Biological and Environmental Research under Award Number DE-SC0014183. Additional support to CEN came from a NIGMS Initiative for Maximizing Student Devel-opment under Award Number R25-GM55036. H.B. and J.G.G. thank the UMBC Eminent Scholar program for catalyzing the ongoing collaboration between the UMBC and UBC groups. GJD is supported through the Royal Society “Ken Murray” Research Professorship.Publisher’s NoteSpringer Nature remains neutral with regard to jurisdictional claims in pub-lished maps and institutional affiliations.Received: 1 December 2017   Accepted: 1 February 2018Page 15 of 16Attia et al. Biotechnol Biofuels  (2018) 11:45 References 1. Parajuli R, Dalgaard T, Jorgensen U, Adamsen APS, Knudsen MT, Birkved M, Gylling M, Schjorring JK. Biorefining in the prevailing energy and materials crisis: a review of sustainable pathways for biorefinery value chains and sustainability assessment methodologies. Renew Sustain Energy Rev. 2015;43:244–63. 2. Tuck CO, Perez E, Horvath IT, Sheldon RA, Poliakoff M. Valorization of biomass: deriving more value from waste. Science. 2012;337:695–9. 3. Horn SJ, Vaaje-Kolstad G, Westereng B, Eijsink VGH. 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