ARTICLE Understanding the Function of Conserved Variations in the Catalytic Loops of Fungal Glycoside Hydrolase Family 12 Andr
e R.L. Dam
asio,1 Marcelo V. Rubio,1,2 Leandro C. Oliveira,1,3 Fernando Segato,1 Bruno A. Dias,1,3 Ana P. Citadini,1 Douglas A. Paix~ao,1 Fabio M. Squina1 1

Laborato´rio Nacional de Ci^encia e Tecnologia do Bioetanol (CTBE), Centro Nacional de Pesquisa em Energia e Materiais (CNPEM), Campinas-SP, Brazil; telephone: þ55 19 3518 3111; fax: þ55 19 35183104; e-mail: [email protected] 2 Instituto de Biologia, Universidade Estadual de Campinas (UNICAMP), Campinas-SP, Brazil 3 Instituto de Bioci^encias, Letras e Ci^encias Exatas, Universidade Estadual Paulista (UNESP), S~ao Jos
e do Rio Preto, SP, Brazil

ABSTRACT: Enzymes that cleave the xyloglucan backbone at unbranched glucose residues have been identified in GH families 5, 7, 12, 16, 44, and 74. Fungi produce enzymes that populate 20 of 22 families that are considered critical for plant biomass deconstruction. We searched for GH12-encoding genes in 27 Eurotiomycetes genomes. After analyzing 50 GH12-related sequences, the conserved variations of the amino acid sequences were examined. Compared to the endoglucanases, the endo-xyloglucanase-associated YSG deletion at the negative subsites of the catalytic cleft with a SST insertion at the reducing end of the substrate-binding crevice is highly conserved. In addition, a highly conserved alanine residue was identified in all xyloglucan-specific enzymes, and this residue is substituted by arginine in more promiscuous glucanases. To understand the basis for the xyloglucan specificity displayed by certain GH12 enzymes, two fungal GH12 endoglucanases were chosen for mutagenesis and functional studies: an endo-xyloglucanase from Aspergillus clavatus (AclaXegA) and an endoglucanase from A. terreus (AtEglD). Comprehensive molecular docking studies and biochemical analyses were performed, revealing that mutations at the entrance of the catalytic cleft in AtEglD result in a wider binding cleft and the alteration of the substrate-cleavage pattern, implying that a trio of residues coordinates the interactions and binding to linear glycans. The loop insertion at the crevice-reducing end of AclaXegA is Andr
e R.L. Dam
asio and Marcelo V. Rubio contributed equally to this work. Correspondence to: F.M. Squina Contract grant sponsor: CNPq Grant numbers: 474022/2011-4; 310177/2011-1 Contract grant sponsor: FAPESP Grant numbers: 2008/58037-9; 2011/02169-4; 2011/13242-7 Contract grant sponsor: FAPESP IC Fellow Contract grant number: 2012/12859-3 Received 15 November 2013; Revision received 24 January 2014; Accepted 27 January 2014 Accepted manuscript online 6 February 2014; Article first published online 27 February 2014 in Wiley Online Library (http://onlinelibrary.wiley.com/doi/10.1002/bit.25209/abstract). DOI 10.1002/bit.25209

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critical for catalytic efficiency to hydrolyze xyloglucan. The understanding of the structural elements governing endo-xyloglucanase activity on linear and branched glucans will facilitate future enzyme modifications with potential applications in industrial biotechnology. Biotechnol. Bioeng. 2014;111: 1494–1505. ß 2014 Wiley Periodicals, Inc. KEYWORDS: fungal endoglucanases; GH12; endoxyloglucanases; xyloglucan specificity

Introduction Xyloglucan (XyG) is the most abundant hemicellulose in the majority of land plants, reaching 20% of the primary cell wall dry weight (Gilbert et al., 2008). XyG is also the primary storage polysaccharide in certain seeds, such as Tamarindus and Hymenaea courbaril (jatob
a) (Buckeridge, 2010). Like cellulose, XyG consists of a linear backbone of b-1,4-glucan linkages but is distinguished by having up to 75% of b-D-Glcp (b-D-glucopyranose) residues that are covalently linked to a-D-Xylp (a-D-xylopyranose) at the O-6 position (Carpita and McCann, 2000). Depending on the source of XyG, a portion of a-D-Xylp residues may be further linked to b-Dgalactopyranose (b-D-Galp) or a-L-arabinofuranose (a-LAraf), and a portion of galactose residues may be extended by a-L-fucopyranose (a-L-Fucp) (Carpita and McCann, 2000). The incubation of XyG with cellulases produces oligosaccharide (XGOs) fingerprints because of an enzyme-specific mode of action combined with the fine structure of the polysaccharide (Buckeridge, 2010; Buckeridge et al., 1992). Enzymes that cleave the XyG backbone at unbranched Glc residues have been identified in GH families 5, 7, 12, 16, 44, and 74. Members of the first five families operate through the canonical double-displacement mechanism of glycosyl ß 2014 Wiley Periodicals, Inc.

transfer, which involves a covalent glycosyl-enzyme intermediate and results in the net retention of the anomeric configuration. Enzymes from GH74 operate by a singledisplacement, anomeric configuration-inverting mechanism involving the direct attack of water on the sugar ring (Gilbert et al., 2008). In addition to endo-acting xyloglucanases, debranching enzymes are needed to complete XyG depolymerization. The XGOs side chains are removed by alpha1,2-L-fucosidases (GH95, EC 3.2.1.63) and glucosidases (GH1, 2, 35, 42, and 43; EC 3.2.1.23), generating xylosylated XGOs (Iglesias et al., 2006), which are further hydrolyzed into monosaccharides by the concerted and sequential actions of xylosidases (GH31; EC 3.2.1.X) and glucosidases (GH1 and GH3; EC 3.2.1.21) (Buckeridge et al., 2000). The basis of endoglucanase specificity to XyG and the ability of certain glucanases to hydrolyze both branched and linear polysaccharides has long been recognized. However, the question posed in 1997 by Vincken et al., that is, “What determines xyloglucanase activity?” has not been fully addressed to date. Conserved sequence variations among the GH12 endo-xyloglucanases and endoglucanases were previously reported (Master et al., 2008). The GH12 endoxyloglucanases have an YSG deletion at the non-reducingend of the catalytic cleft (loop 1) and a SST insertion at the reducing end of the substrate-binding crevice (loop 2). In addition, there is a highly conserved alanine residue in loop 2 in all endo-xyloglucanases that is replaced by arginine in more promiscuous GH12 glucanases. This study aimed to elucidate the structural role of conserved sequence variations in loops at the entry and exit of the catalytic cleft that define fungal GH12 xyloglucan-specific enzymes. The GH12-encoding genes derived from 27 Eurotiomycetes genomes support the hypothesis that the conserved sequence deletion and insertion variations are structural determinants. To understand the basis of the xyloglucan specificity displayed by certain GH12 enzymes, two fungal GH12 endoglucanases were chosen for mutagenesis and functional studies: the endo-xyloglucanase derived from Aspergillus clavatus (AclaXegA) and the promiscuous endoglucanase from A. terreus (AtEglD). Molecular docking studies and biochemical analysis of the GH12 mutants comprehensively described the role of the loops at the xyloglucan/b-glucan interaction site in the catalytic cleft and during hydrolysis.

selected and grown in 5-mL LB-ampicillin broth, the plasmids were purified (Sambrook et al., 1988) and digested with NotI/XbaI, and the insert size was verified by 1% agarosegel electrophoresis (Sambrook et al., 1988). Plasmids with the correct insert size were fully sequenced, and clones with the correct DNA sequence were used for the transformation of Aspergillus nidulans strain A773 (pyrG89; wA3; pyroA4), as previously described (Segato et al., 2012). Next, 107–108 spores/mL were inoculated in liquid minimal medium supplemented with 5% maltose, distributed onto dishes and incubated without shaking at 37 C for 2– 3 days. The mycelial mat was lifted with a spatula and discarded, and the medium was collected by filtration, centrifuged at 10,000g for 10 min prior to concentration by ultra-filtration (10,000-Da cutoff, Millipore, Billerica, MA), quantified by the Bradford method (Bradford, 1976), validated for purity by SDS–PAGE (Shapiro et al., 1967) and used in biochemical studies. Site-Directed Mutagenesis Site-directed mutagenesis was carried out by the standard PCR-based method using AtEglD and AclaXegA parental genes as templates. The amplified fragments were used as templates for the overlap-extension PCR technique (Heckman and Pease, 2007) to fuse the two genes in a single ORF. The fused fragment was digested with the restriction enzymes NotI and XbaI and cloned into pEXPYR (Segato et al., 2012). Protein Purification The target proteins were purified in two steps. The concentrated and dialyzed protein samples were applied to an ion-exchange Resource Q column equilibrated with 20 mM sodium phosphate buffer, pH 7.4, and the proteins were eluted with a linear 0-to-1 M sodium chloride gradient (Äkta Purifier, GE, Little Chalfont, UK). Fractions active on beta-glucan or xyloglucan were collected and loaded onto a Superdex G-75 (10 mm  30 mm) gel-filtration column and equilibrated with 50 mM ammonium acetate buffer, pH 5.0, and eluted fractions showing enzymatic activity were analyzed by SDS–PAGE. Single-band fractions were combined, concentrated and used for further biochemical analysis. The flow rate used for both chromatographic steps was 0.5 mL min1. Purified fractions were validated by SDS–PAGE.

Materials and Methods Enzymatic Properties Cloning and Expression of AtEglD and AclaXegA PCR-amplified gene fragments were digested with NotI and XbaI, after which they were isolated by excising a thin slice from a 0.8%-agarose electrophoresis gel and purified with a QIAquick Gel Extraction kit (Qiagen, Venlo, NL). The fragments were then ligated into NotI/XbaI-digested pEXPYR plasmid with T4-fast ligase (Promega, Fitchburg, WI) and transformed into Caþ-competent DH5a Escherichia coli (Promega). Random ampicillin-resistant colonies were

Reducing sugars were determined using 3,5-dinitrosalicylic acid (DNS) and monitored colorimetrically at 540 nm (Miller, 1959) using an Infinite1 200 PRO microplate reader (TECAN, Mannedorf, CH). One unit of enzyme was defined as the quantity of enzyme needed to release reducing sugars at rate of 1 mmol/min under standard conditions. The standard assay was conducted for 10 min in 50 mM McIlvaine glycineadded buffer at pH 5.5, 50 C, with substrates at 2.0 mg mL1 and 1 mg of purified enzymes.

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The optimal pH and temperature were determined for AtEglD and AclaXegA activities using barley b-glucan and xyloglucan from tamarind (XyG), respectively, as substrates under standard conditions. The assays for substrate specificity were evaluated in various substrates, including arabinan from the sugar beet, debranched arabinan, linear arabinan, rye arabinoxylan, larch arabinogalactan, galactomanan, XyG, oat-spelt xylan, wheat arabinoxylan, barley b-glucan, carboxymethylcellulose (CMC) and xylan from beechwood. The polysaccharides were purchased from Sigma Aldrich (St. Louis, MO) or Megazyme, Co. (Wicklow, IE). The kinetic parameters were estimated for all enzymes from initial rates at 11 substrate concentrations of 1–14.4 and 1–10.8 mg mL1 for b-glucan and XyG, respectively. The assays were carried out under standard conditions to assess Vmax, Km, and Kcat. Capillary Electrophoresis (CE) Oligosaccharides (Megazyme) were derivatized with 8aminopyreno-1,3,6-trisulfonic acid (APTS) by reductive amination (Naran et al., 2007). Enzymatic hydrolysis of labeled substrates was performed at 50 C. To analyze the cleavage patterns, capillary-zone electrophoresis (CZE) of substrate-breakdown products was performed using a P/ACE MQD instrument (Beckman Coulter, Pasadena, CA) equipped with a laser-induced fluorescence detector. A fusedsilica capillary (TSP 050375, Polymicro Technologies, Phoenix, AZ) with an internal diameter of 50 mm and total length 31 cm was used as separation column for oligosaccharides. The electrophoresis conditions were as follows: 30 kV/70–100 mA at 20 C using sodium phosphate buffer (40 mM, pH 2.5). Because of the small volumes of capillary electrophoresis combined with the small variations in buffer strength, retention times vary slightly when comparing separate electrophoresis runs. Circular Dichroism Spectra of far-UV circular dichroism (CD) were taken on a JASCO J-810 spectropolarimeter (Jasco, Inc., Tokyo, Japan) equipped with a Peltier temperature-control unit using a wavelength range of 195–240 nm, and a 0.1-cm-path quartz cuvet, and the solvent spectra were subtracted in all experiments to eliminate background effects. CD spectra were the average of eight accumulations taken using a scanning speed of 100 nm min1, a spectral bandwidth of 1 nm, and a response time of 0.5 s. The protein concentrations were 0.2 mg mL1 in 50 mM sodium phosphate buffer, pH 7.4. Thermal denaturation was characterized by measuring the changes in ellipticity at 218.5 nm induced by an increase in temperature from 20 to 100 C at 1 C min1 (Cota et al., 2011). Homology Molecular Modeling and Analysis of Wild and Mutant Types The initial structure was modeled using homologous enzymes extracted from the protein data bank (PDB). The

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atomic coordinates from Aspergillus niger endoglucanase (PDB id: 1KS5) (Khademi et al., 2002) and Aspergillus aculeatus xyloglucanase (PDB id: 3VL8) (Yoshizawa et al., 2012) were used as templates to generate structural models for AtEglD by restraint-based modeling, as implemented in the MODELLER program (Sali and Blundell, 1993) in the HHpred Server (Soding et al., 2005). The mutants were constructed by removing the target regions, connecting the extremities where the region was removed and applying the Dunbrack rotamer libraries (Dunbrack, 2002) in the UCSF Chimera package, version 1.7 (Pettersen et al., 2004). Next, the conformation energy was minimized using the Chimera interface with the AMBER ff12SB force field employed to 5000 steepest descent steps and 5000 conjugate gradient  steps, both of which had a size of 0.02 A. The changes in the dynamic behavior of all enzymes were analyzed using Normal Modes (NMA) (Hollup et al., 2005). The NMA approach models the residues using beads in the alpha-carbon position. The application uses a harmonic potential to define the residue interactions and analyze the low frequency normal modes. The fluctuations are calculated using the atomic displacement (Di), given by d2 Di ¼ Pni 2  1 di where n is the total number of residues in the protein and di is the component of the eigenvector corresponding to the i residue. The fluctuations are normalized to have a maximum peak at 1.0. Molecular Docking Cellohexaose and xyloglucan oligosaccharide structures were built using GLYCAM04 (Basma et al., 2001; Kirschner and Woods, 2001a,b). Docking was performed using the procedure described by Sagermann (Sagermann and Matthews, 2002) through UCSF Chimera and UCSF dock (Lang et al., 2009; Moustakas et al., 2006; Pettersen et al., 2004). The enzymes and ligands were initially prepared using the Prep Dock tool in the Chimera interface (AMBER ss12SB force field for proteins and AM1-BCC force fields for other cases). The cavity of the binding site was suggested by the docking program and is consistent with the results of recent work (Gloster et al., 2007). The spatial conformations of the enzymes were aligned, and the grid box was defined similarly for all of the conformations. Three hundred docking runs were performed for each substrate. The ligand, receptor,  ligand orientations and overlap bins were set to 0.2 A, and the distance tolerance for matching between the atoms and the  receptor was set to 0.75 A. The conformations obtained from the docking runs were analyzed for the distance between the central oxygen (O4) of the ligand and the C-alpha position of the residue ALA:102 in AclaXegA and AclaXegADSST and between the central oxygen and the C-alpha position of the residue ALA:97 in AtEglD and

AtEglDDYSG. These distances were organized in histograms  using bins of 2 A, emphasizing the differences in substrate accommodation and displacement.

Results and Discussion The Majority of the Eurotiomycetes Genomes Contain a Single Gene for Endo-Xyloglucanase The comparative analysis of the filamentous fungal genomes revealed a large number of CAZy genes, ranging from 171 to 285 per genome. For example, Trichoderma reesei contains approximately 200 GH enzymes and A. niger encodes approximately 250 GHs (Jovanovic et al., 2009). According to Jovanovic et al. (2009), although 115 GH families are recognized by the CAZY database, only 22 families contain enzymes critical for plant biomass deconstruction. Fungi produce enzymes in 20 of these families, covering all activities needed for the efficient conversion of natural biomass (Jovanovic et al., 2009). The class Eurotiomycetes (Ascomycota, Pezizomycotina) is a monophyletic group comprising two major clades of very different ascomycetous fungi: (i) the subclass Eurotiomycetidae; and (ii) the subclass Chaetothyriomycetidae. Eurotiomycetidae includes producers of toxic and useful secondary metabolites, fermentation agents used to make food products and enzymes, xerophiles and psychrophiles, and the important genetic model A. nidulans (Geiser et al., 2006). We searched for GH12-encoding genes in 27 Eurotiomycetes genomes using the JGI MycoCosm tool (Grigoriev et al., 2012), revealing 50 related GH12 sequences. The genetic model Neurospora crassa (Znameroski and Glass, 2013)

and the model for biomass deconstruction, T. reesei, were chosen as non-Eurotiomycetes external groups. Apart from A. brasiliensis and A. flavus, all analyzed Eurotiomycetes have one copy of a GH12 endo-xyloglucanase-encoding gene (Supplementary Table SI). A. carbonarius, A. glaucus, A. nidulans, and A. zonatus have no copies of GH12 endoglucanase-encoding genes. Interesting, N. crassa carries no GH12 enzymes. Moreover, we did not find any GH12 endo-xyloglucanase-encoding genes in the T. reesei genome (Supplementary Table SI). The phylogenetic tree of Aspergilli GH12 enzymes revealed two well-defined clades and a common ancestor of GH12 endoglucanases and endo-xyloglucanases (Fig. 1). Sequences in the same clade exhibited high similarity, ranging from 52% to 70% amino acid identity. Eurotiomycetes GH12 Endo-Xyloglucanases (3.2.1.151) Have Conserved Deletions and Insertions Compared to GH12 Endoglucanases After careful analysis of 50 GH12-related sequences from 27 Eurotiomycetes, the conserved variations of endo-xyloglucanase and endoglucanase amino acid sequences were noteworthy. Compared to the endoglucanase amino acid sequences, the endo-xyloglucanases have two highly conserved variations located in loop 1 and loop 2 (Fig. 2A), and these variations can be observed across all the analyzed Eurotiomycetes (Supplementary Fig. S1). The predicted molecular models for AclaXegA and AtEglD exhibited a typical b-jelly roll tertiary structure described for GH12 (Fig. 2) (Gloster et al., 2007). These two structures share the same topology, with mainly b-strands in the concave region (Fig. 2B and C). These models of AclaXegA

Figure 1.

Phylogenetic tree of fungal endoglucanases from glycoside hydrolase, family 12. All of these sequences are derived from eukaryotes (except the root sequence) and previously characterized enzymes from the Carbohydrate-Active Enzymes (CAZy) database (www.cazy.org). The scale bar indicates branch length. For the phylogenetic analysis, the amino acid sequences were aligned using ClustalX 1.83 software. The phylogenetic tree was built using Mega 4 software. The highlighted symbols represent the GH12 enzymes studied in this work: AclaXegA (ACLA_029940) and AtEglD (ATEG_09894).

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Figure 2. Comparison of the AclaXegA and AtEglD structures. A: Alignment of the primary sequences of AclaXegA and AtEglD. Endoglucanase 3 from Trichoderma harzianum (ThEG3) was adopted as a reference. B and C: The b-jellyroll structure of AclaXegA (template PDB: 3VL8; C-score 1.76) and AtEglD (template PDB: 1KS5A; C-score 1.88). The highlighted blue (Panel B) and orange (Panel C) regions indicate the deletions. The asterisk indicates an arginine that is highly conserved in endoglucanases and substituted by alanine in xyloglucanases. D: Close-up of the substrate-binding cleft of fungal glycoside GH12. The side chains of some of the most important residues for AclaXegA (blue) and AtEglD (red) are drawn. The surfaces on the right side highlight the same residues in the cleft. The asterisks denote the deletion (YSG) and insertion (SST) in AclaXegA.

and AtEglD allowed the depiction of conserved amino acid variations among GH12-related sequences. Specifically, the GH12 endo-xyloglucanases show a deletion event at the nonreducing end of the catalytic cleft (loop 1) and an insertion event at the reducing end of the substrate-binding crevice (loop 2). Accordingly, AclaXegA has an YSG deletion in the loop between B5 and B6 (loop 1) (Fig. 2B) and a SST insertion in the loop between B6 and B8 (loop 2) (Fig. 2C). In addition to these conserved variations, there is a highly conserved alanine (A) residue in loop 2 in all GH12 endoxyloglucanases (Supplementary Figs. S1 and S2A) that is replaced by arginine (AtEglD arginine 123 (R123)) in more promiscuous GH12 glucanases (Fig. 2A). The YSG AclaXegA deletion shortens loop 1 at the nonreducing end of the catalytic cleft, subtracting an aromatic interaction described as critical for cellopentaose substrate binding in T. harzianum GH12 endoglucanase (Prates et al., 2013). In addition, loop 2 is longer in fungal GH12 endo-xyloglucanases than in endoglucanases. This insertion

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in AclaXegA loop 2 is adjacent to the so-called “cord” region (P131, I132), which is conserved in all analyzed fungal GH12 amino acid sequences (Supplementary Fig. S1). The cord region was previously described and contributes with amino acid residues to the substrate-binding cleft that are likely involved in binding the reducing end of the substrate (Sandgren et al., 2001). As shown in Figure 2D, the residues of the catalytic cleft are highly conserved between fungal GH12s. Previous elegant structural studies of fungal and bacterial GH12 enzymes have provided the basis to define substrate binding to the cord region, as well as the core residues of the cleft that are likely to be involved in binding the reducing end of the substrate (þ1 and þ2 subsites) (Gloster et al., 2007; Sandgren et al., 2001). All these characteristics can indicate the substrate binding properties of xyloglucan-specific enzymes relative to more promiscuous endoglucanases, as well as the inability of fungal GH12 endo-xyloglucanases to hydrolyze unbranched substrates (Sandgren et al., 2001; Master et al., 2008).

AclaXegA and AtEglD Are Typical Fungal Glycoside Hydrolases From Family 12 To understand the structural basis of xyloglucan specificity displayed by certain GH12 enzymes, we chose two fungal GH12 endoglucanases to conduct mutagenesis and functional studies: the endoglucanase-encoding gene from A. terreus (ATEG_09894/AtEglD) and the endo-xyloglucanaseencoding gene from A. clavatus (ACLA_029940/AclaXegA). The pattern of hydrolysis of the unbranched XyG backbone by fungal GH12 enzymes has been examined previously (Damasio et al., 2012; Master et al., 2008; Song et al., 2013), including comprehensive studies on substrate recognition, as in the description of the A. niger xyloglucanspecific GH12 (AnXEG12A), which prefers xyloglucanoligosaccharides containing more than six glucose units, and a study of the importance of the xylose substitution at the 3 and þ1 positions (Powlowski et al., 2009). AclaXegA is a strict xyloglucan-specific enzyme that hydrolyzes xyloglucan at unbranched glucose residues (Fig. 3A and C). Moreover, AtEglD is a promiscuous endoglucanase that preferentially hydrolyzes undecorated glucan backbones (b-glucan) over xyloglucan (Fig. 3A). AtEglD and AclaXegA share the same mode of operation, releasing XXXG, XLXG, and XLLG from xyloglucan as the final products (data not shown). The hydrolysis of b-glucan by AtEglD released cellotetraose, cellotriose, and cellobiose as major products (Fig. 3B). Protein Secretion and Folding To ensure proper post-translational modifications the genes described in this work were cloned into the pEXPYR shuttle vector and were transformed in A. nidulans to reach high levels of target protein secretion (Segato et al., 2012). The target proteins (AclaXegA, AclaXegADSST, AtEglD, AtEglDDYSG, AtEglDR123A) were purified by two chromatographic-purification steps. The secondary structures were evaluated by circular-dichroism (CD) (Supplementary Fig. S2), and the data were analyzed using the DichroMatch database (Supplementary Table SII). AtEglD and AclaXegA showed a predominance of b-strand secondary structures. Despite AtEglDDYSG, the mutants (AtEglDR123A and AclaXegADSST) had similar CD profiles compared to the parental enzymes. Although AtEglDDYSG exhibited a change on the CD profile, the b-strand was also the predominant secondary structure. The Dichromatch analyses suggest a possible structural rearrangement with b-strand loss, and the gain of irregular structure for AtEglDDYSG (Supplementary Table SII). The SST Deletion in AclaXegA Did Not Affect the Enzymes Affinity for the Substrate But Did Reduce the Catalytic Efficiency The characterization by Normal-Mode Analysis (NMA) revealed discrete fluctuation changes in the deleted loop

Figure 3. Substrate specificity and mode of operation of AclaXegA and AtEglD on polysaccharides. A: AclaXegA was highly specific to xyloglucan hydrolysis. B: Capillary electrophoresis of APTS-labeled oligosaccharides after the hydrolysis of bglucan by AtEglD and of xyloglucan from tamarind by AclaXegA (C). APTS-labeled glucose (C1), cellotetraose (C4), cellopentaose (C5) and cellohexaose (C6). The APTSlabeled xyloglucan oligosaccharides (XXXG, XLXG, and XLLG). XyG, xyloglucan from tamarind; CMC, carboxymethylcellulose. The segments of the xyloglucan polymer are named based on a one-letter unambiguous system. Unsubstituted D-Glcp is designated as G; the a-D-Xylp-(1 ! 6)-b-D-Glcp residue is designated as X; the b-D-Galp-(1 ! 2)a-D-Xylp-(1 ! 6)-b-D-Glcp is designated as L. The assay was carried out using AtEglD or AclaXegA at 0.086 mM and the substrate at 10 mg/mL in 50 mM ammonium acetate buffer, pH 5.5 for 5 h at 50 C.

region (S133, S134, and T135) (Fig. 4A and B). These data corroborate the CD results, as the denaturation midpoint (Tm) was exactly the same for AclaXegA and AclaXegADSST (Fig. 4C). Additionally, almost no changes were observed in protein secondary structures (Supplementary Fig. S2). The mode of operation of AclaXegADSST was the same as that observed for the parental enzyme, releasing XXXG, XLXG, and XLLG as major products from xyloglucan (Fig. 4D) as well as the pH and temperature optimum (Supplementary Fig. S3). The best fit of the substrate in the catalytic cleft was reached after three hundred molecular docking attempts; however, the substrate positioning in the catalytic crevice was exactly the same in all attempts for the parental AclaXegA (Fig. 6A).

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Figure 4. Dynamic behavior of AclaXegA and AclaXegADSST evaluated by normal mode analysis (NMA). A: The mobility profile along the primary sequence of AclaXegA, expressed via the fluctuations normalized of the carbon alpha atoms using the 7–12 low-frequency normal modes. The black box highlights the mutated region. B: The protein regions that the displacement changed during the analysis are highlighted by red balls. The asterisk correlates the mutated region in the graph to that in the predicted structure. C: Thermal denaturation curve at pH 7.4. The thermal denaturation curve was obtained by monitoring at 217.6 nm. D: Capillary electrophoresis of APTS-labeled oligosaccharides after hydrolysis of xyloglucan from tamarind (XyG) by AclaXegA and AclaXegADSST for 2 h. The xyloglucan oligosaccharide nomenclature is described in Figure 3.

The docking analysis for AclaXegADSST revealed that the substrate fit poorly in the catalytic cleft due to a loss of interaction at the cord region (Fig. 6B), mainly at the þ2 and þ3 subsites. Conversely, the interactions in the negative subsites were maintained, highlighting the importance of W8 and W23, which are critical to substrate placement in the crevice. According to previous studies, the a-D-Xylp substitution at the 3 position is necessary for efficient hydrolytic activity, and substrate occupation of the hydrophobic 4 position improves substrate binding 30-fold (Powlowski et al., 2009). The Km for AclaXegADSST on XyG was unchanged, unlike the Vmax and the turnover number (Kcat), which were significantly reduced. Thus, the catalytic efficiency of AclaXegADSST for xyloglucan hydrolysis decreased approximately eightfold (Table I). This result is in agreement with the docking data. The SST deletion changed the enzymesubstrate binding interactions, most notably at the AclaXegA positive subsites, resulting in substrate misfitting and

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instability at the catalytic cleft and changes in hydrolytic performance. According to our findings, after the loop 2 truncation in AclaXegA (AclaXegADSST), the cord region residues, in particular Pro129 and Ile130 which are likely to form the bottom of the þ2 and þ3 subsites (Sandgren et al., 2005), were displaced (Fig. 6A and B). These findings underline the role of positive subsites in substrate interaction with the catalytic crevice, along with the xylosyl substitution of XyG at the þ1 site. Indeed, the xylosyl substitution at the þ1 position is essential for substrate binding and hydrolysis, and more than one positive subsite is required for efficient hydrolysis (Powlowski et al., 2009). The histogram (Fig. 7A) illustrates the distribution of docking populations obtained for AclaXegA and AclaXegADSST. The parental enzyme showed a narrow distribution compared to AclaXegADSST. These differences corroborate the variations in substrate accommodation at the negative subsites, as well as the inferior substrate-conversion

Table I. Wild-type and mutant kinetic parameters. Protein

Substrate

Km (mg/mL)

Vmax (mmol product/min/mM enzyme)

Kcat (s1)

Kcat/Km

AtEglD

b-Glucan XyGa b-Glucan XyG b-Glucan XyG XyG XyG

5.39  1.3 4.74  2.9 5.04  1.1 5.51  1.8 5.41  1.2 1.53  0.14 2.38  0.82 1.89  0.49

463  45 219  25 240  20 115  16 10.6  1.1 1.39  0.037 218  24 22.5  1.5

77.1 36.5 40.0 19.2 1.74 0.230 36.3 3.70

14.3 7.70 7.94 3.48 0.322 0.150 15.3 1.95

AtEglDR123A AtEglDDYSG AclaXegA AclaXegADSST

ND, not detected. Xyloglucan from tamarind.

a

rate of AclaXegADSST, which is caused by the spatial restrictions needed to adjust the substrate in the catalytic cleft. The YSG Deletion in AtEglD Altered Catalysis and Reduced Thermal Stability The trio of residues that contribute important hydrophobic interactions (Y7, W22, Y111; using the residue numbering of AtEglD) at the non-reducing end of the catalytic cleft is highly conserved in fungal GH12 endoglucanases (EC 3.2.1.4). Y7,

W28, and Y111 are conserved in 100%, 100%, and 40% of the endoglucanase sequences, respectively, in 27 Eurotiomycetes genomes analyzed in this study. While Y7 can be substituted by tryptophan in some endo-xyloglucanases; Y111 is absent in all endo-xyloglucanases and substituted by serine in 60% of the endoglucanases analyzed in this study (Supplementary Fig. S1). The characterization of AtEglD and mutants using NMA indicated that several protein regions displayed differences in mobility after YSG deletion (AtEglDDYSG), most notably in the truncated loop 1 region (Fig. 5A and B). This result is

Figure 5. Evaluation of the dynamic behavior of AtEglD, AtEglDDYSG, and AtEglDR123A using NMA. A: The NMA evaluation of the fluctuations, normalized as described in Figure 4A. The black box highlights the YSG deletion region. B: The protein regions changed by the displacement during AtEglDDYSG analysis are highlighted with red balls. The asterisks indicate the YSG deletion in the graph and in the predicted structure. C: Thermal-denaturation curve at pH 7.4. The thermal-denaturation curve was obtained by monitoring at 217.6 nm. Capillary electrophoresis of APTS-labeled oligosaccharides after hydrolysis of xyloglucan from tamarind (D) and APTS-labeled cellohexaose for 2 h (E).

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Figure 6. Molecular docking analysis. AclaXegA (A) and AclaXegADSST (B) in complex with XXXG/XXX. Red: conserved aromatic residues (W8 and W23) at the nonreducing end of the catalytic cleft; Light blue: residues of the cord region (P131, I132); Yellow: the deleted residues (SST) in AclaXegADSST. AtEglD (C) and AtEglDDYSG (D) in complex with cellohexaose (C6). Red: conserved aromatic residues (Y7, W22, and Y111) at the non-reducing end of the catalytic cleft. The Y111 residue was deleted in AtEglDDYSG. Figure 7. Histograms generated by molecular docking. Histograms of the distance distributions of the 300 molecular docking experiments run for wild-type and mutants to AclaXegA (A) and AtEglD (B). in accordance with CD analysis, as higher-mobility AtEglDD YSG can explain the reduction of mutant Tm at over 10 (Fig. 5C). The pH and temperature optima were unaltered after YSG deletion (Supplementary Fig. S4). The catalytic efficiency of AtEglDDYSG in the xyloglucan and b-glucan degradation was significantly reduced (Table I). The Y111 residue is absent in the AtEglDDYSG loop 1, as previously mentioned. Thus, the lack of this key residue in AtEglDDYSG was expected to have meaningful implications for substrate–enzyme interactions. After the AtEglDDYSG deletion, cellohexaose was misfitted at the catalytic cleft (Fig. 6C and D). The mode of operation of AtEglDDYSG was unchanged for xyloglucan hydrolysis (Fig. 5D). In the other hand, the mode of operation on b-glucan hydrolysis was changed, producing cellotriose as a major product after cellohexaose hydrolysis (Fig. 5E). The conformations obtained after docking runs showed a single distribution for the parental enzyme and cellohexaose fit into the binding site (Fig. 7B). Conversely, the cellohexaose swings away the catalytic cleft of AtEglDDYSG, and the correct alignment between the catalytic triad and the

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substrate is not reached most of the time (Fig. 7B) due to the YSG deletion at the cleft entrance (Fig. 6D). Our results provide a biochemical basis for the MD simulations reported by Prates et al. (2013). According to Prates et al., an inspection of the trajectories for a GH12 endoglucanase from T. harzianum (ThEG3) cellotetraose and cellopentaose models reveals that after approximately 10 ns, the substrate swings away from residues Y7 and W23 and is not oriented along the crevice but remains connected to the enzyme by the Y112 hydrophobic contact. After 20–30 ns, the substrate fits back into the crevice in a conformation that resembles that of ThEG3 (Prates et al., 2013). The AtEglDDYSG model also indicates that the catalyticcleft volume is increased (Fig. 6D), thus validating the previous insights of Powlowski et al. (2009); the more open conformation of the binding cleft predicted for AnXEG12A (endo-xyloglucanase) compared to HgGH12 (endoglucanase) may provide interactions with linear polymeric substrates. Conversely, the narrow substrate-binding cleft of HgGH12 compared to AnXEG12A could explain the

reduced activity of HgGH12 on branched polysaccharides (Powlowski et al., 2009). Here, the wider binding cleft observed for AclaXegA was correlated with the lack of activity on b-glucan. Similarly, the narrower cleft of AtEglD was related to a lower efficiency in xyloglucan hydrolysis. Accordingly, the YSG deletion caused further enlargement of the volume of the crevice, followed by the reduction of b-glucan-hydrolysis capacity (Table I). The R123A Mutation in AtEglD Led to Vibration Mode Variations and Reduced the Catalytic Efficiency The R123 in AtEglD is a highly conserved residue in fungal GH12 endoglucanases and is replaced by alanine in all the GH12 fungal endo-xyloglucanases analyzed here (Supplementary Fig. S1). Although the mode of operation and the pH and temperature optima were unchanged (Fig. 5D and E and Supplementary Fig. S4), the catalytic efficiency of AtEglDR123A was reduced twofold for b-glucan and xyloglucan hydrolysis (Table I). Prates et al. (2013) suggested, based on MD simulations, that the interaction of the R124 in ThEG3 (equivalent to R123 in AtEglD) with I128 is critical for B9 strand stabilization. Accordingly, this stabilization was missed in the AtEglDR123A mutant.

Figure 8. Oligosaccharide hydrolysis by AtEglD and AtEglDDYSG. Capillary electrophoresis after hydrolysis of APTS-labeled cellopentaose (C5) (A) and cellohexaose (C6) (B) for 12 h. The wider arrows indicate the preferred cleavage site.

The dynamic behavior, evaluated by NMA, revealed that several regions displayed differences in the vibration mode (Fig. 5A), such as the residues in the B9 strand and its adjacent loop (residues from 138 to 147, Fig. 2). In accordance with the CD analysis, this higher mobility compared to the parental enzyme reduced the mutant protein Tm by over 10 . The AtEglD R123 is located in loop 2 and is also adjacent to the cord region previously discussed for the AclaXegADSST mutant. Accordingly, the AtEglDR123A mutation also altered substrate–enzyme interactions at the positive subsites, reducing catalytic efficiency but not affecting the enzyme’s affinity for the substrate (Table I). The Loop 1 Extension in Fungal GH12 Endoglucanases Governs Linear Glucan Hydrolysis The loop 1 extension (YSG) at the non-reducing end of the catalytic cleft is essential to the interactions at the 4 and 3 subsites and hydrolysis of linear b-glucan. The CE analysis of the AtEglD mode of operation indicated that cellobiose was the major product after APTS-labeled cellopentaose hydrolysis and that cellobiose and cellotetraose were produced from APTS-labeled cellohexaose, suggesting the key role of the 3 and 4 GH12 subsites for the hydrolysis of linear glucans (Fig. 8). Moreover, AtEglD did not hydrolyze cellotetraose (data not shown), as this substrate can only interact with the subsite region from 2 to þ2. Indeed, the inefficient substrate interaction at the 3 subsite by AtEglDDYSG resulted in reduced catalytic efficiency; after a 12-h hydrolysis, cellopentaose (C5) was only partially hydrolyzed (Fig. 8A). Consequently, AtEglDDYSG interactions at the negative subsites were drastically affected (Figs. 6D and 7B), causing substrate misfitting in the catalytic crevice and a significant reduction in the efficiency of linear b-glucan degradation (Table I). Furthermore, the absence of YSG extension in AclaXegA resulted in a wider crevice than that of AtEglD (Fig. 6). To summarize, our reports describe the role of structural determinants in fungal GH12 enzymes. All fungal GH12 endo-xyloglucanases have conserved variations at loop 1 and loop 2 compared to GH12 endoglucanase counterparts. The conserved deletion (YSG) shortens loop 1 at the nonreducing end of the catalytic cleft, contributing to a more open binding cleft compared to fungal GH12 endoglucanases. Conversely, the GH12 endo-xyloglucanase SST insertion at loop 2, which is located in the positive subsites, is essential for the correct binding of xyloglucan and thus for the catalytic efficiency of endo-xyloglucanase. Lastly, the arginine residue (R123 for AtEglD) that is conserved in all the fungal GH12 endoglucanases is important for loop 2 stabilization; loop 2, in turn, coordinates the substrate interactions at the positive subsites. These conclusions not only contribute to a better understanding of the specificity of fungal GH12 endoxyloglucanases but can also help to direct efforts for the rational design of site-directed mutagenesis and targeting

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enzymes for specific applications. Deciphering the molecular determinants of catalysis is of high relevance to biotechnology with possible applications for the bioproduction of addedvalue chemicals. This work was financially supported by grants from CNPq (474022/ 2011-4 and 310177/2011-1) and FAPESP (2008/58037-9). ARLD (2011/02169-4) and LCO (2011/13242-7) are FAPESP postdoctoral fellows. MVR is a FAPESP IC fellow (2012/12859-3). Authors’ Contributions: A.R.L.D. designed the study, designed the site-directed mutagenesis, performed heterologous expression of the parental enzymes in A. nidulans, analyzed the results and wrote the manuscript. M.V.R. performed the mutant cloning and expression, purification, biochemical characterizations, and capillary electrophoresis. L.C.O. conducted the normal-mode analysis, molecular docking and computational data evaluation and participated in manuscript preparation. A.P.C. participated in circular-dichroism spectra measurement. B.A.D. and F.S. performed the biochemical characterizations of the parental enzymes and capillary electrophoresis and participated in manuscript preparation. D.A.P. built the phylogenetic tree and analyzed the data. F.M.S. revised the manuscript and coordinated the study. All authors have read and approved the final manuscript.

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