Novel Endotype Xanthanase from Xanthan-DegradingMicrobacterium sp. Strain XT11

Fan Yang,a He Li,a Jie Sun,a Xiaoyu Guo,a Xinyu Zhang,a Min Tao,a Xiaoyi Chen,a Xianzhen Lia

aSchool of Biological Engineering, Dalian Polytechnic University, Ganjingziqu, Dalian, People’s Republic China

ABSTRACT Under general aqueous conditions, xanthan appears in an ordered con-formation, which makes its backbone largely resistant to degradation by known cel-lulases. Therefore, the xanthan degradation mechanism is still unclear because of thelack of an efficient hydrolase. Here, we report the catalytic properties of MiXen, axanthan-degrading enzyme identified from the genus Microbacterium. MiXen is a952-amino-acid protein that is unique to strain XT11. Both the sequence and struc-tural features suggested that MiXen belongs to a new branch of the GH9 family andhas a multimodular structure in which a catalytic (�/�)6 barrel is flanked by anN-terminal Ig-like domain and by a C-terminal domain that has very few homo-logues in sequence databases and functions as a carbohydrate-binding module(CBM). Based on circular dichroism, shear-dependent viscosity, and reducing sugarand gel permeation chromatography analysis, we demonstrated that recombinantMiXen efficiently and randomly cleaved glucosidic bonds within the highly orderedxanthan substrate. A MiXen mutant free of the C-terminal CBM domain partially lostits xanthan-hydrolyzing ability because of decreased affinity toward xanthan, indicat-ing the CBM domain assisted MiXen in hydrolyzing highly ordered xanthan via rec-ognizing and binding to the substrate. Furthermore, side chain substituents and theterminal mannosyl residue significantly influenced the activity of MiXen via the for-mation of barriers to enzymolysis. Overall, the results of this study provide insightinto the hydrolysis mechanism and enzymatic properties of a novel endotype xan-thanase that will benefit future applications.

IMPORTANCE This work characterized a novel endotype xanthanase, MiXen, andelucidated that the C-terminal carbohydrate-binding module of MiXen could drasti-cally enhance the hydrolysis activity of the enzyme toward highly ordered xanthan.Both the sequence and structural analysis demonstrated that the catalytic domainand carbohydrate-binding module of MiXen belong to the novel branch of the GH9family and CBMs, respectively. This xanthan cleaver can help further reveal the enzy-molysis mechanism of xanthan and provide an efficient tool for the production ofmolecular modified xanthan with new physicochemical and physiological functions.

KEYWORDS conformation, endoxanthanase, xanthan backbone, xanthan hydrolysis

Xanthan, an anionic heteropolysaccharide with a molecular mass ranging from1 � 106 to 7 � 106 Da, is produced by the plant-pathogenic bacterium Xanthomo-

nas campestris pv. campestris (1). Xanthan contains a cellulosic chain as a backbone anda linear mannosyl-glucuronyl-mannosyl trisaccharide as a side chain attached at the C-3position on the alternate glucosyl residue of the main chain (2, 3). The inner andterminal mannosyl residues of the side chains are often acetylated and pyruvylated,respectively, depending on both the xanthan-producing strains and the culture con-ditions (2). The complex structure endows xanthan with superior rheological properties,such as pseudoplasticity, high viscosity, and tolerance toward a wide range of pHs and

Citation Yang F, Li H, Sun J, Guo X, Zhang X,Tao M, Chen X, Li X. 2019. Novel endotypexanthanase from xanthan-degradingMicrobacterium sp. strain XT11. Appl EnvironMicrobiol 85:e01800-18. https://doi.org/10.1128/AEM.01800-18.

Editor Haruyuki Atomi, Kyoto University

Copyright © 2019 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Xianzhen Li,xianzhen@mail.com.

H.L. and J.S. contributed equally to this work.

Received 31 July 2018Accepted 27 October 2018

Accepted manuscript posted online 9November 2018Published

BIODEGRADATION

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temperatures, leading to its widespread application as the thickener and stabilizer inthe food, pharmaceutical, and oil industries (4, 5).

To date, molecular modified xanthan with novel physicochemical and physiologicalfunctions has been sought for use in new applications. Although genetically engi-neered Xanthomonas mutants have been reported to produce variant xanthan prod-ucts, their production levels are far from those needed for practical application (1, 6, 7).Preparation of such modified xanthan by chemical or physical degradation of thexanthan backbone is difficult because of the complex structure of the polymer (4). Inaddition, chemical or physical modification of xanthan is nonspecific and will also causedegradation of the xanthan side chains (8). Thus, enzyme-assisted methods thatspecifically attack the xanthan backbone while leaving the side chains intact seem tobe a promising method for preparation of molecularly designed xanthan.

In aqueous solution, native xanthan exists in a double-stranded helical conformationalong with an order-disorder transition (8). Previous studies have demonstrated thatxanthan in a highly disordered conformation can be hydrolyzed by cellulases (9, 10).However, the accessibility of xanthan by enzymes could be dramatically reduced whenthe ordered fraction in xanthan increases (8, 11). It should be noted that the confor-mation of xanthan is highly dependent on changes in temperature, ionic strength, andmolecular composition of xanthan (12, 13), particularly with respect to the presence ofacetyl and/or pyruvate groups (14, 15), and completely disordered xanthan can only beobtained in solutions with very low ionic strength and at a high temperature (8), whichis difficult to achieve under industrial conditions. Furthermore, considering that enzymeactivities can also be affected by temperature and ionic strength, low enzyme activitymay occur under conditions in which xanthan appears in a completely disorderedconformation.

Currently, little is known about the hydrolysis mechanism of individual enzymes onthe xanthan backbone and enzyme resistance of highly ordered xanthan remains abottleneck for the production of modified xanthan. In this study, an endotype xantha-nase, MiXen, from the xanthan-degrading bacterium Microbacterium sp. strain XT11,was cloned and heterologously expressed. The sequence/structure homology, bio-chemical characteristics, and product profiles of MiXen were studied, revealing novelhydrolysis properties concerning the high substrate specificity and hydrolysis efficiencyof MiXen against xanthan with a highly ordered conformation.

RESULTSSequence analysis and homology modeling of MiXen reveals a multidomain

structure. Previously, an aerobic Gram-positive bacterium, Microbacterium sp. XT11,with the ability to degrade xanthan, was isolated from a soil sample collected from theDalian Botanical Garden in China (16), and the complete genome sequence of strainXT11 was determined and analyzed (17). Within the genomic sequence of Micro-bacterium sp. XT11, the open reading frame encoding MiXen (GenBank accessionnumber ALX66163.1) is 2,856 bp in length and has a GC content of 67.3%. Thepredicted molecular mass of MiXen is 101.1 kDa, and its calculated isoelectric point is4.4. SignalP (version 4.1) analysis showed that the predicted signal peptide of MiXenwas composed of 33 amino acid residues (Met1 to Ala33). Functional domains of MiXenwere predicted by the InterProScan web server. MiXen consisted of a cellulaseN-terminal Ig-like domain (Ala34 to Glu124), a putative family 9 glycoside hydrolase(GH9) catalytic domain (Asp125 to Ser580), and a suspected carbohydrate-bindingmodule (CBM) at the C terminus (Arg700 to Glu848).

Previous reports confirmed that structures of the catalytic domains of GH family 9enzymes are characterized by an (�/�)6-barrel fold with three acidic active-site residues(two aspartate residues and one glutamate residue) (18). The C terminus-free MiXenshowed highest identity scores with three characterized GH9 enzymes, including GH9endoglucanase LC-CelG from an uncultured bacterium (accession number AHL27900,24% identity), cellobiohydrolase CbhA from Clostridium thermocellum (PDB entry 1RQ5,21% identity), and endoglucanase AaCel9A from Alicyclobacillus acidocaldarius (acces-

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sion number ACV59481, 19% identity). Sequence alignment of the C terminus-freeMiXen with LC-CelG, CbhA, and AaCel9A revealed three conserved active-site residues(Asp192, Asp195, and Glu568) in the catalytic domain of MiXen (Fig. 1a). Secondarystructural analysis predicted that the catalytic domain of MiXen belongs to a typical(�/�)6-barrel fold, which contains 12 long �-helices forming the central (�/�)6-barreland 6 antiparallel strands forming two �-sheets (Fig. 1a). Taken together, these primaryand secondary structural features suggest that MiXen belongs to the GH9 family.

Homology modeling was also performed to further predict the typical GH9 structureof MiXen. The top three identified structural analogs of MiXen, LC-CelG (PDB entry3X17), CbhA (PDB entry 1RQ5), and AaCel9A (PDB entry 3GZK), were used to constructmodels of MiXen without the C-terminal domain. Structures 3X17, 1RQ5, and 3GZKhave a bond length root mean square deviation of around 0.77 Å, 2.79 Å, and 1.89 Å forthe models, respectively, which would reflect the similarity of the protein cores. Figure1b depicts a hypothetical structure of MiXen without the C-terminal domain. Overall,68% of the input sequence was modeled at �65% coverage (structural analog, PDBentry 3X17), mainly covering a globular catalytic domain (54.2% of amino acid residuesin �-helices) and a cellulase N-terminal Ig-like domain (69.1% amino acid residues in�-strands and coils). The structural superposition of the modeled MiXen with 3X17revealed that the steric configuration of the MiXen catalytic domain is nearly identicalto that of 3X17 with a typical GH9 structure (Fig. 1c). However, the low sequenceidentity of the MiXen catalytic domain to 3X17, 1RQ5, and 3GZK suggested that MiXenrepresents a novel branch of the GH9 family.

Recombinant expression and characterization of MiXen. The gene encodingMiXen free of the N-terminal signal peptide was amplified from the genomic DNA ofMicrobacterium sp. XT11. The DNA products were gel purified and cloned into theexpression vector. To investigate the function of the C-terminal domain of MiXen, tworecombinants, MiXen-CD (MiXen mutant without the C-terminal domain) and MiXen-CT(the C-terminal domain of MiXen), were constructed, after which the proteins wereheterologously expressed in E. coli BL21(DE3). Following the purification, proteins wereexamined to confirm the purity and molecular mass by SDS-PAGE. The purified solubleproteins MiXen, MiXen-CD, and MiXen-CT had a purity of �90% and migrated as singlebands with approximate molecular masses of 100 kDa, 80 kDa, and 25 kDa, respectively(see Fig. S1 in the supplemental material).

To determine the apparent temperature/pH optima and stability of the enzymes,carboxymethyl cellulose (CMC) rather than xanthan was chosen as the model substrate.This was because (i) the conformation of xanthan is susceptible to temperature and pHand (ii) the backbone of xanthan is cellulose-like. MiXen presented its highest activityat 40°C to 45°C and retained more than 70% of its residual activity after incubation for30 min at temperatures ranging from 20°C to 45°C (Fig. 2a). The optimal pH value forMiXen was 7.5 to 8.0 (Fig. 2b). The pH stability profile showed that MiXen could retainmore than 80% of its residual activity after incubation in buffers with pHs ranging from6 to 9 for 2 h (Fig. 2b). Notably, the maximum activity was obtained when theconcentration of the NaH2PO4-Na2HPO4 buffer was 10 mM. When CMC degradation wasconducted in deionized water, dramatically decreasing activity was observed (Fig. 2c).Therefore, the following enzymolysis experiments, including the determination of kineticparameters, were conducted in 10 mM NaH2PO4-Na2HPO4 buffer at 40°C and pH 7.5.

MiXen could attack xanthan with a highly ordered conformation. As a key factorthat affects the degree of xanthan enzymolysis, the disordered fraction (�) of thepurified native xanthan sample was measured by circular dichroism. When xanthan wassuspended in 10 mM NaH2PO4-Na2HPO4 buffer (pH 7.5) at 40°C, the disordered fractionwas 0, which reflected that xanthan was folded in a highly ordered conformation underthe enzyme assay conditions. As shown in Table 1, after 12 h of incubation with MiXen,the disordered fraction of xanthan digests increased to 0.425 � 0.018. A smaller disor-dered fraction of 0.193 � 0.023 was observed with another monomodular xanthanase,MiGH from Microbacterium sp. XT11 (19), indicating that MiXen degraded more highly

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FIG 1 Sequence and structure properties of MiXen. (a) Alignment of amino acid sequences of MiXen, LC-CelG, AaCel9A, and CbhA. Theaccession numbers were ALX66163 for MiXen, AHL27900 for LC-CelG, ACV59481 for AaCel9A, and 1RQ5 for CbhA. The amino acidsequence of MiXen without a putative signal peptide and a C-terminal domain (residues 34 to 580) and the corresponding regions of the

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ordered xanthan than MiGH. Moreover, the xanthan substrate maintained a highlyordered conformation after incubation with the commercial cellulases or xanthan lyase,which suggested that cellulases and xanthan lyase could not act on the xanthanmolecule.

The shear-dependent viscosity of xanthan digests was recorded in Fig. 3a. After 12 hof incubation with different enzymes, shear-thinning was observed for all xanthandigests being investigated. The MiXen-digested xanthan showed a low-shear-rate(0.015 s�1) viscosity of about 75.0 Pa·s, which was lower than those observed byMiGH-digested xanthan (83.8 Pa·s), xanthan lyase-digested xanthan (93.6 Pa·s),cellulase-digested xanthan (94.0 Pa·s), and native xanthan (96.8 Pa·s). The decreasedviscosity indicated that the backbone of highly ordered xanthan could be effectively

FIG 1 Legend (Continued)other three proteins are shown. The amino acid residues, which are conserved in all four proteins, are denoted with white letters andhighlighted in black. The amino acid residues, which are conserved in two or three different proteins, are highlighted in gray. The rangesof the secondary structures of MiXen (�a–�g strands for the Ig-like domain, �1–�6 strands and �1–�13 helices for the catalytic domain)are shown above its sequence based on the predicted crystal structure of MiXen. The conserved residues that form the catalytic site(Asp192, Asp195, and Glu568) and substrate binding site (Tyr199, His511, and Arg513) are indicated by asterisks and plus signs,respectively, above the MiXen sequence. (b) Dimensional cartoon structure of the C-terminus-free MiXen was predicted using the I-TASSERprotein structure homology-modeling on-line server (http://zhanglab.ccmb.med.umich.edu/I-TASSER/). The following colors were used toindicate distinct domains in MiXen: cellulase N-terminal Ig-like domain, violet; glycoside hydrolase family 9, green. (c) A structural overlayof the modeled MiXen with LC-CelG (PDB entry 3X17) was made to allow visual comparison. MiXen, green; LC-CelG, blue.

FIG 2 Biochemical characteristics of MiXen. (a) The temperature optimum (�) was measured by incubating 5 g/liter carboxymethylcellulose (CMC) with 0.15 mg/ml MiXen for 20 min at the indicated temperatures. To determine the thermostability (�), the remainingenzyme activity was measured at 40°C after incubation at the indicated temperatures for 30 min. (b) The pH optimum (�) was determinedby measuring the activity at the indicated pH values. For the pH stability (�), enzyme activity was measured at 40°C after incubation atthe indicated pH for 2 h. (c) The effects of the NaH2PO4-Na2HPO4 buffer concentration on enzyme activities.

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cleaved by MiXen, whereas the cellulases and xanthan lyase could not react with thexanthan backbone.

The specific activities of MiXen and other selected enzymes against highly orderedxanthan were then investigated. As shown in Fig. 3b and Table 1, cellulases exhibitedthe highest activity toward CMC, while there was no significant difference between thespecific activity of MiXen and MiGH toward CMC. However, when xanthan was used asthe substrate, the MiXen activity was remarkably higher than the MiGH activity, whileno activity was detected when cellulases or xanthan lyase was added. Moreover, the

TABLE 1 Hydrolysis properties of different enzymes on xanthan backbone

Enzyme(s)Disordered fraction ofxanthan digestsa (�) Sp act (U/g) Km (g/liter) kcat (min�1) kcat/Km (liters/g·min)

MiXen 0.425 � 0.018 15.70 � 0.98 0.65 � 0.02 9.80 � 0.20 15.08 � 0.38MiXen-CD 0.167 � 0.013 5.12 � 0.59 1.18 � 0.02 3.37 � 0.05 2.86 � 0.05MiXen-CT 0 0MiGH 0.193 � 0.023 9.20 � 1.22 0.94 � 0.01 7.29 � 0.48 7.75 � 0.62Xanthan lyase 0 0Cellulases 0 0.47 � 0.25aThe disordered fractions of xanthan digests (�) were determined after incubation with enzymes at 40°C and pH 7.5 for 12 h.

FIG 3 Hydrolysis performance of different enzymes against highly ordered xanthan. (a) Viscosity of native xanthan as a function of shearrate after being hydrolyzed by different enzymes. Xanthan degradation was conducted at 40°C and pH 7.5 for 12 h. The final concentrationof the enzyme was 1.0 mg/ml. Xanthan incubated with bovine serum albumin was used as a negative control. All samples were preparedin duplicate, and the averages are presented. (b) Specific activities of enzymes toward different substrates. The final concentration ofMiXen, MiXen-CD, MiXen-CT, MiGH, or xanthan lyase was 0.1 mg/ml. The concentration of mixed cellulases was 0.1 mg/ml for xanthansubstrates and 0.01 mg/ml for CMC. Xanthan and CMC (5 g/liter) were incubated with enzymes at 40°C and pH 7.5 for 20 min, respectively.The data represent the averages � standard deviations from at least three independent samples. *, P � 0.05. (c) GPC elution patterns ofxanthan after incubation with different enzymes at 40°C for 12 h. The final concentration of the enzyme was 1.0 mg/ml.

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catalytic efficiency (kcat/Km) of MiXen was 2-fold higher than that of MiGH, and the Km

value of MiXen was significantly lower than that of MiGH, suggesting a higher substratespecificity of MiXen on xanthan with a highly ordered conformation.

The molecular mass distributions of xanthan digests produced by excessiveamounts of enzymes were also detected by gel permeation chromatography (GPC) (Fig.3c). After 12 h of incubation with MiXen, the xanthan enzymolysis products weredivided into two fractions, nondegraded high-molecular-mass (HMW) xanthan (reten-tion time [Rt] �18 to 21 min) and some intermediate degradation products (Rt � 21 to24 min). Compared with MiXen, MiGH degraded HMW xanthan into intermediatedegradation products with a wider range of molecular masses. Consistent with theobservations in the viscosity and enzyme activity assays, cellulases failed to cleave thebackbone of highly ordered xanthan.

MiXen exhibits endotype xanthanase activity. MiXen resembles (24% sequenceidentity) the catalytic part of a GH9 from uncultured bacterium (PDB entry 3X17), whichhas been shown to be an endotype glucanase that exerts significant activities againstCMC but no activity toward p-nitrophenyl cellobioside (18). Gel permeation chroma-tography analysis also provided information about the xanthan-degrading pattern ofMiXen with time (Fig. 4). As the incubation time increased, more HMW xanthan wasconverted into intermediate degradation products. After 6 h of incubation with1 mg/ml MiXen, no more degradation of the high-molecular-mass xanthan was ob-served, and no completely degraded low-molecular-mass (LMW) xanthan digests (Rt �

28 to 33 min) were detected. Moreover, further increases in enzyme concentration didnot lead to the production of additional xanthan digests (data not shown). The GPCresult suggests that MiXen could not cleave at the termini of the xanthan backbone. Tofurther assess the randomness of cleavage of the xanthan backbone by MiXen, therelationship between the production of reducing groups and increase in fluidity wasmonitored. As shown in Fig. 5, there was a proportionate and steep increase in fluiditywith increases in reducing sugar, which demonstrated that MiXen could randomlycleave glucosidic bonds within the xanthan backbone to produce intermediate xanthandigests, suggesting an endotype hydrolytic attack of MiXen.

The C-terminal domain helps MiXen bind to highly ordered xanthan. Comparedwith the wild-type protein, the truncated protein free of CBM (MiXen-CD) showeddramatically decreased xanthan-degrading ability. The higher low-shear-rate viscosity(89.8 Pa·s) and smaller disordered fraction (0.167 � 0.013) of MiXen-CD-digested xan-than suggested a weakened interaction with xanthan (Fig. 3a and Table 1). The specificactivity of MiXen-CD on xanthan did not differ significantly from that on CMC and wasnearly 2-fold lower than the MiXen activity on xanthan (Fig. 3b). As shown in Fig. 3c, asimilar elution pattern was observed when xanthan was digested by truncated MiXen.

FIG 4 GPC elution patterns of xanthan after incubation with 1.0 mg/ml MiXen for different times.

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Compared to MiXen, MiXen-CD showed slightly weaker hydrolysis performance viadegrading less xanthan into intermediate degradation products. Michaelis-Mentenconstants were also determined. As shown in Table 1, the catalytic efficiency (kcat/Km)of MiXen was 5-fold higher than that of MiXen-CD. In addition, the Km value of thetruncated protein was significantly higher than that of MiXen, representing a decreasedaffinity for xanthan. Taken together, these data indicate that the C-terminal domainplays a key role in xanthan hydrolysis by enhancing the substrate affinity of theenzymes.

The low-shear-rate viscosity and disordered fraction of MiXen-CT-digested xanthanwere 94.5 Pa·s and 0, respectively (Fig. 3a and Table 1). No activity was observed whenxanthan was incubated with MiXen-CT. All of the results indicate that the C-terminaldomain of MiXen could not cleave the backbone of xanthan. The surface plasmonresonance (SPR) was then designed to measure the interaction between MiXen-CT andthe xanthan substrate. As shown in Fig. 6, the C-terminal domain of MiXen exhibited astrong affinity toward xanthan with a low dissociation constant (KD) value at 0.651 �g/ml, indicating that the C-terminal domain assists MiXen in hydrolyzing highly orderedxanthan via adsorption on the substrate.

Modification of xanthan side chain affects the hydrolysis activity of MiXen bychanging the xanthan secondary structure. To determine the structural transition ofxanthan upon changes in the xanthan side chain, xanthan samples with different side

FIG 5 Relationship between fluidity and production of reducing sugar groups during hydrolysis ofxanthan by MiXen.

FIG 6 SPR analysis of the interaction of MiXen-CT and native xanthan. Different traces correspond toincreasing xanthan concentrations. The dissociation constant (KD) values were calculated with a one-sitesteady-state binding model.

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chain modifications, including pyruvate-free xanthan (PFX), acetyl-free xanthan (AFX),acetyl- and pyruvate-free xanthan (APFX), and terminal mannosyl residue-free xanthan(TMFX), were prepared. Tables 2 and 3 provide an overview of chemical compositionsof different xanthan samples and the corresponding fraction of disordered conforma-tion (�). Except for TMFX, the disordered fraction of which was detected as0.100 � 0.015, all other xanthan samples showed no disordered fraction under enzy-molysis conditions. After 12 h of incubation with MiXen, the disordered fraction of allxanthan digests increased. The TMFX digests had the largest disordered fraction,0.537 � 0.021, probably because of the partially disordered conformation of the initialsubstrate. The NX digests also gained a relatively larger disordered fraction(0.425 � 0.018) than the AFX, PFX, and APFX digests. These results suggest that, afterbeing incubated with MiXen, more disordered molecules were produced from NX andTMFX than from AFX, PFX, and APFX.

Because the effects of acetyl and/or pyruvate elimination on the xanthan structurecould not be significantly observed through order-disorder transitions, the variations insecondary structure among different modified xanthans were investigated directlyusing atomic force microscopy (AFM). As shown in Fig. 7a, purified native xanthan (NX)exposed visible branches, giving it a tree-like structure, and the mean height of the NXsample was estimated to be 0.66 � 0.05 nm (Fig. 7f), suggesting a double-strandedsecondary structure. In contrast to those of NX, strands of PFX, AFX, and APFX becamemore agglomerated and complex and exhibited many sharp kinks (Fig. 7b to d). Higherstrand heights of the three samples were also observed (PFX, 0.84 � 0.07 nm; AFX,0.83 � 0.09 nm; APFX, 0.85 � 0.10 nm) (Fig. 7f), which was probably due to the highabundance of locally folded polymers. Taken together, these changes would makexanthan strands more enzyme resistant. Unlike the other samples, terminal mannosylresidue-free xanthan (TMFX) exposed more random-coil structures, with the lowestmean height being 0.48 � 0.07 nm (Fig. 7e and f), suggesting more single-strandedpolymers, which is consistent with a previous report (20). The mean height of xanthansamples after incubation with MiXen was then calculated. As shown in Fig. 7f, com-pared with that of the undigested samples, the mean height of all degraded xanthansamples decreased, indicating that some locally folded polymers or other complexsecondary structures have been transformed into disordered structures. Compared with

TABLE 2 Summarized properties of xanthan with different modifications

Xanthan samplea

Content (wt/wt, %)Disordered fractionb

(�)Acetyl Pyruvate

NX 13.2 9.6 0AFX 0 8.2 0PFX 10.2 5.0 0APFX 0 5.7 0TMFX 11.8 6.4 0.100 � 0.015aXanthan samples include native xanthan (NX), pyruvate-free xanthan (PFX), acetyl-free xanthan (AFX), acetyl-and pyruvate-free xanthan (APFX), and terminal mannosyl residue-free xanthan (TMFX).

bThe fractions of disordered conformation (�) were determined at 40°C and pH 7.5.

TABLE 3 Summarized characteristics of MiXen against different xanthan substrates

Characteristic

Value for xanthan substrateb

NX AFX PFX APFX TMFX

Disordered fraction of xanthan digestsa (�) 0.425 � 0.018 0.183 � 0.025 0.242 � 0.013 0.250 � 0.016 0.537 � 0.021Sp act (U/g) 15.7 � 0.98 13.0 � 0.54 11.2 � 0.35 10.2 � 0.27 19.4 � 1.05Km (g/liter) 0.65 � 0.02 1.05 � 0.03 0.85 � 0.06 1.00 � 0.10 0.52 � 0.01kcat (min�1) 9.80 � 0.20 9.59 � 0.67 8.56 � 0.14 6.24 � 0.17 12.87 � 0.21kcat/Km (liters/g·min) 15.08 � 0.38 9.13 � 0.78 10.07 � 0.20 6.24 � 0.21 24.75 � 0.49aThe disordered fractions of xanthan digests (�) were determined after incubation with MiXen at 40°C and pH 7.5 for 12 h.bXanthan substrates include native xanthan (NX), pyruvate-free xanthan (PFX), acetyl-free xanthan (AFX), acetyl- and pyruvate-free xanthan (APFX), and terminalmannosyl residue-free xanthan (TMFX).

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those of digests produced from AFX, PFX, and APFX, TMFX and NX digests showedrelatively lower mean heights of 0.36 � 0.08 nm and 0.48 � 0.08 nm, respectively,suggesting more disordered molecules formed. These results were in accordance withthose observed in the circular dichroism assay.

The specific activities and Michaelis-Menten constants were measured after theenzyme MiXen was incubated with different xanthan modifications. Table 3 summarizes

FIG 7 AFM analysis of different xanthan samples. All xanthan samples were suspended in 10 mM NaH2PO4-Na2HPO4 buffer (pH7.5) at a final concentration of 1 g/liter. (a to e) AFM topography images of native xanthan (NX), pyruvate-free xanthan (PFX),acetyl-free xanthan (AFX), acetyl- and pyruvate-free xanthan (APFX), and terminal mannosyl residue free xanthan samples(TMFX), respectively. Figure contrast was automatically adjusted with XEP software (versions 1.7.70.3; Park Systems, SouthKorea) to enable better visualization. (f) Boxplots of the measured heights of the different xanthan samples before (blank) andafter (gray) incubation with 1.0 mg/ml MiXen at 40°C for 12 h. The sample sizes were 15 (NX), 15 (PFX), 20 (AFX), 18 (APFX),and 17 (TMFX).

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the hydrolysis parameters of MiXen. The activities of MiXen on AFX, PFX, and APFXgreatly decreased compared to that on the NX sample, and the simultaneous absenceof acetyl and pyruvate groups caused the maximum decline of enzyme activity. Themaximum MiXen activity was obtained in the TMFX-hydrolyzing system. The Michaelis-Menten constants of MiXen were also determined for different xanthan samples. The Km

values of MiXen on AFX (Km � 1.05 g/liter), PFX (Km � 0.85 g/liter), and APFX (Km � 1.00g/liter) were significantly higher than that of MiXen on NX (Km � 0.65 g/liter), indicatingdecreased substrate affinities. However, a slightly lower Km value was calculated forTMFX (Km � 0.52 g/liter). The catalytic efficiencies (kcat/Km) of MiXen on AFX, PFX, andAPFX were lower than that of MiXen on NX, and the highest kcat/Km value wascalculated when TMFX was used as the substrate. The enzymatic parameters of MiXentoward different xanthan samples were consistent with the xanthan structural proper-ties observed using circular dichroism and AFM.

DISCUSSION

Similar to cellulose, the full degradation of which depends on synergistic interac-tions of glycoside hydrolases (21), exhaustive degradation of the cellulose-like back-bone of xanthan should be the result of the collective activity of glycoside hydrolases.To date, many studies have investigated xanthan degradation using mixed cellulasesystems prepared from cellulolytic microorganisms (8, 10, 11). However, few studieshave been conducted to elucidate the hydrolysis properties of individual enzymes onxanthan. A xanthanase in Microbacterium sp. strain XT11, MiXen, is presumed to be akey xanthan-degrading enzyme because (i) the gene encoding MiXen is located in axanthan-degrading gene cluster in strain XT11 (17) and (ii) sequence and structuralfeatures suggest that MiXen belongs to the glycoside hydrolase family 9 (Fig. 1), mostmembers of which have been characterized as cellulases. Interestingly, because of itslow sequence and structural homologies to previously identified enzymes (�25%),MiXen may represent a novel branch of the GH9 family. Additionally, the secondarystructural prediction for MiXen revealed a multidomain structure, with a function-unknown C-terminal module that might confer some unique hydrolysis properties toMiXen.

Circular dichroism revealed that, in a solution with high ionic strength (Na� con-centration, �10 mM) at 40°C, native xanthan exists in a highly ordered conformation(� � 0) (Table 2), which was previously confirmed to be completely resistant to thecommercial cellulases from SEAB (France) (11). In the present study, MiXen showedhigher specific activity and affinity toward highly ordered xanthan than other selectedenzymes and exhibited efficient hydrolysis performance by greatly increasing thedisordered fraction and decreasing the viscosity of xanthan (Fig. 3 and Table 1),suggesting that such highly ordered xanthan exhibited less resistance to MiXen. Inaddition to having cellulase activity, many GH9 cellulases have been reported to displayside activities on related polysaccharides, such as glycans (22), xylans (23), or xyloglu-cans (24), but they most prefer soluble (CMC and cellodextrins) or insoluble (Avicel)cellulose (25). Unlike other known GH9 cellulases, the specific activity of MiXen onxanthan was much higher than that on CMC (Fig. 3), whereas the Km value againstxanthan was significantly lower than that against CMC, indicating that xanthan is thefavorite substrate for MiXen. A considerable portion of the high-molecular-mass xan-than remained after 36 h of incubation with an excessive amount of MiXen, which wasprobably because of the multiple stranded networks of xanthan helices (26). Never-theless, MiXen should still be considered a good cleaver compared to the otherenzymes selected in this study and previously reported cellulase mixtures that couldnot attack highly ordered xanthan (� � 0.02), even after 48 h of incubation (8).

Based on the investigations described above, we investigated what makes MiXenactive specifically on highly ordered xanthan and how it is distinct from the other GH9cellulases. Earlier studies of modular cellulases focused on bacterial enzymes possess-ing cellulose-binding CBM, such as family 17, family 28, and family 2a (27–29). More-over, previous biochemical studies showed that CBM increases the amount of enzyme

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bound on cellulose, indicating that the main role of CBM is to increase the affinity tocellulose (30, 31). Similar to a large proportion of known GH9 enzymes, MiXen pos-sesses an ancillary C-terminal module in addition to the catalytic domain and an Ig-likedomain, which was confirmed to be a carbohydrate-binding module by SPR analysis(Fig. 6). However, it should be noted that both sequence and structural similarities ofCBM in MiXen were lower (�10%) than those of other known cellulase CBMs, indicatingthe C-terminal domain of MiXen is a representative of a novel branch of CBMs.Consistent with previous demonstrations that removal of CBM significantly reduced theactivity of cellulases toward insoluble substrates (32, 33), the CBM domain-truncatedMiXen-CD lost its substrate specificity for xanthan while also exhibiting dramaticallydecreased degradation activity, substrate affinity, and catalytic efficiency (Fig. 3 andTable 1). In addition, the CBM domain showed no activity against xanthan, indicatingthat CBM is simply a helper for substrate recognition. Taken together, these resultsconfirmed the hypothesis that the CBM domain assists MiXen in hydrolyzing highlyordered xanthan by recognizing and binding to the substrate.

It is known that the final degree of enzymolysis could be influenced by thesecondary xanthan structure (8), and that changes in xanthan structure depend ondifferent factors, including temperature, pH, ionic strength, and modifications of thexanthan side chain (20, 34, 35). In this study, xanthan degradation was conducted underset temperature, pH, and ionic strength; thus, the effect of xanthan side chain modi-fication on MiXen activity via changes in xanthan structure was further investigated.The removal of pyruvate and/or acetyl groups from xanthan, along with the increasingelectrostatic interactions and/or rigidity, led to higher strand heights and the formationof high entanglement points and locally folded polymers (Fig. 7), which is in agreementwith the results of previous studies (20, 26, 36). These changes indicated thatsubstituent-free xanthan samples were in a highly ordered state and formed barriers toenzymolysis, explaining the dramatically decreased hydrolysis capacity of MiXen onAFX, PFX, and APFX observed in this study. Notably, compared to native xanthan, whichhas side chains that revolve around the main-chain skeleton by reverse winding (37),TMFX had a slightly larger disordered fraction (� � 0.1) and more flexible structure,which provided more accessibility to enzymatic degradation. Thus, MiXen showed thehighest hydrolysis ability against TMFX.

Conclusions. MiXen is a xanthan hydrolase obtained from a xanthan-degradingbacterium, Microbacterium sp. XT11, and probably belongs to a novel branch of theGH9 family. This enzyme can efficiently and randomly cleave glucosidic bonds withinthe backbone of highly ordered xanthan, and the novel C-terminal carbohydrate-binding module of MiXen plays a key role in xanthan recognition and binding.Moreover, this study demonstrated that the modification of xanthan side chainsinfluences the hydrolysis activity of MiXen by changing xanthan secondary structures.

MATERIALS AND METHODSStrains and culture conditions. Xanthan-degrading strain Microbacterium sp. XT11 (deposited at the

China Center for Type Culture Collection [CCTCC AB2016011]) was isolated by the laboratory of X. Li (16)and used in this study for the cloning of endoxanthanase-encoding genes. Escherichia coli DH5� wasused in all cloning experiments. Escherichia coli BL21(DE3) was capable of the expression of endoxan-thanase.

Microbacterium sp. XT11 was cultivated at 30°C in xanthan medium (3 g/liter xanthan, 0.5 g/literglucose, and 3 g/liter yeast extract dissolved in mineral salt solution, pH 7.0). The mineral salt solutioncontained (0.05 g/liter K2HPO4, 0.8 g/liter NaCl, 0.025 g/liter MgSO4·7H2O, and 0.70 g/liter KNO3). E. colistrains were grown at 37°C in Luria-Bertani (LB) medium (10 g/liter tryptone, 5.0 g/liter yeast extract, and10 g/liter sodium chloride, pH 7.0) supplemented with 100 mg/liter ampicillin or kanamycin if necessary.

Sequence analysis and modeling of enzymes. Protein annotation was conducted using theCarbohydrate Active Enzyme ANnotation web server (dbCAN; http://cys.bios.niu.edu/dbCAN2/blast.php)(38). Functional domains of the protein were predicted using the InterProScan web server (http://www.ebi.ac.uk/interpro/search/sequence-search) (39). To obtain reference data for MiXen, the online auto-mated protein structure homology-modeling server I-TASSER (http://zhanglab.ccmb.med.umich.edu/I-TASSER/) was employed to predict the protein structure (40). The amino acid sequence of MiXen(GenBank accession number ALX66163.1) without the signal peptide (amino acids 1 to 33) was used formodeling.

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Cloning, expression, and purification of endoxanthanase constructs. The coding gene of MiXenwithout the signal peptide was amplified by PCR using the genomic DNA template extracted fromMicrobacterium sp. XT11. The forward primer was 5=-CCCGAATTCGCCACCATCGACAAGGTCACGG-3=(EcoRI restriction site underlined), and the reverse primer was 5=CGCAAGCTTTCAGCCCACAGCGACT-3=(HindIII restriction site underlined). To construct the C-terminal domain of MiXen and the mutant missingthe C terminus, plasmid pET-MiXen was used as the PCR template. The DNA fragment encoding the Nterminus and the catalytic domain was deleted using the forward primer 5=-CCCGAATTCGCGTCGGGCAGGTGACCAAC-3=, and the resulting mutant was named MiXen-CT. The DNA fragment encoding theC-terminal domain was deleted using the reverse primer 5=-CGCAAGCTTGGCATGGTTGTTGTACCAATTCG-3=,and the resulting mutant was named MiXen-CD. To amplify the gene encoding another previouslyreported monomodular xanthanase from Microbacterium sp. XT11 (19), the amino acid sequence was firstdetermined by liquid chromatography-tandem mass spectrometry (LC-MS/MS), which revealed that thisxanthanase is identical to a superfamily glycoside hydrolase, MiGH (GenBank accession numberWP_067195711), in the genome annotation database of strain XT11 (see Fig. S2 in the supplementalmaterial). The primers used to amplify the MiGH coding genes were 5=-CCCGAATTCATGCGCCCCACCATCG-3= (EcoRI restriction site underlined) and 5=-CGCAAGCTTTCAGACGACGGTGTTCTG-3= (HindIIIrestriction site underlined).

All of the PCR products were digested with EcoRI and HindIII and then ligated into the EcoRI- andHindIII-cut pET28a vector. A 6� His tag sequence was added to the 5= terminus of DNA products fordownstream protein purification. The generated recombinant plasmids pET-MiXen, pET-MiXen-CD, pET-MiXen-CT, and pET-MiGH were transformed into electrocompetent E. coli BL21(DE3) by electroporationat 2.5 kV, 25 �F, and 200 in a 0.2-cm-gap electroporation cuvette using a Bio-Rad Gene Pulser. Therecombinants were selected on LB plates containing 100 mg/liter kanamycin.

For the expression of MiXen, MiXen-CD, MiXen-CT, and MiGH, E. coli transformants were grown at37°C in LB medium supplemented with 30 mg/liter kanamycin until the optical density at 600 nm (OD600)reached 0.5. For induction, isopropyl-�-D-thiogalactopyranoside (IPTG) was added at a final concentrationof 0.5 mM, after which the cells were further cultured at 16°C for 12 h. Cells next were harvested bycentrifugation, and the cell pellet was suspended and sonicated 20 times on ice for 10 s with 30-sintervals between cycles using an ultrasonic processor (Branson Digital Sonifier; Branson, Inc., USA).Following removal of cell debris by centrifugation, the supernatant was ready for subsequent purifica-tion. His-tagged proteins were purified using a nickel-nitrilotriacetic acid (Ni-NTA) purification system aspreviously described (41), and the N-terminal His tag was then cleaved using thrombin and removedusing Ni-NTA Sepharose. The proteins were subsequently fractionated on a Superdex 75 gel filtration column(GE Healthcare, Uppsala, Sweden) to remove residual thrombin. The purity of the fractions was assessed bySDS-PAGE. The purified protein concentration was measured using a Bio-Rad protein assay kit.

Enzyme assay. The specific activity of the enzyme was determined by incubating 5 g/liter xanthanor carboxymethyl cellulose (CMC) with appropriately diluted enzymes in 10 mM NaH2PO4-Na2HPO4

buffer (pH 7.5). The final concentration of MiXen, MiXen-CD, MiXen-CT, MiGH, or xanthan lyase (42) usedin the assay was 0.1 mg/ml. The concentrations of mixed cellulases (Sigma) from Aspergillus niger were0.1 mg/ml for xanthan substrates and 0.01 mg/ml for CMC. The reaction was conducted at 40°C for20 min and then terminated by boiling the mixture for 5 min. Under these conditions, the formation ofreducing-end sugars was proportional to the reaction time, and less than 5% of the substrate wasconsumed. The liberation of reducing-end sugars was assayed by the 2,2’-bicinchoninate method (43).One unit of enzyme activity was defined as the amount of enzyme that produced 1 �mol reducing-endsugar per min. All assays were conducted in triplicate, and the mean values were presented.

To assess the optimum temperature, enzyme activity was measured at temperatures ranging from20°C to 70°C according to the standard procedure described above. The thermostability of the enzymewas determined by assaying the residual enzyme activity after the enzyme was incubated at varioustemperatures (20°C to 60°C) for 30 min. The optimal pH for activity was assayed by measuring theenzyme activity at 40°C in different buffers, ranging from pH 2 to 10. To measure the pH stability, activityassays were performed using enzyme samples that were preincubated at 4°C for 2 h in different buffers,ranging from pH 2 to 10. Buffers were prepared at a 10 mM concentration at 0.5 pH intervals, includingglycine-HCl buffer (pH 2.0 to 4.0), acetic acid-sodium acetate buffer (pH 4.0 to 6.0), NaH2PO4-Na2HPO4

buffer (pH 6.0 to 8.0), and glycine-NaOH buffer (pH 8.0 to 10.0). The effect of buffer concentration onenzyme activity was examined by determining activity in different concentrations of NaH2PO4-Na2HPO4

buffer. All reactions to optimize activity were conducted using a final enzyme concentration of 0.15 mg/ml.Under the optional conditions, kinetic parameters (Km and Vmax) were determined by incubating the

enzyme with different concentrations of xanthan solution (0.1, 0.25, 0.5, 0.75, 1.0, 1.5, 2.0, 2.5, 4.0, and 5.0g/liter). According to the time course for the formation of reducing-end sugars, the initial velocity wascalculated as the slope of a tangent at the origin. Km and Vmax were calculated using the Michaelis-Menten equation.

Hydrolysis of xanthan substrates (5 g/liter) was accomplished in 10 mM NaH2PO4-Na2HPO4 buffercontaining an excess of enzyme at 1 mg/ml at 40°C for the specified times.

All assays were conducted in triplicate, and the mean values are presented.Surface plasmon resonance. SPR measurement was performed on a BIAcore T200 instrument

with a sensor chip with dextran matrix that was precoated with streptavidin (GE Healthcare). Thechip was activated with 420-s injections of 0.05 M N-hydroxysuccinimide and 0.2 M ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride. MiXen-CT diluted with running buffer was injectedand captured onto the channel surface to achieve an immobilization level of about 2,000 RU (responseunits). For the binding assay, the reaction temperature was controlled at 25 � 0.01°C and the flow rate

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was set to 30 �l/min. The response obtained from the detection channel (xanthan) was normalized bysubtracting the signal simultaneously acquired from the control channel, which could eliminate non-specific binding and buffer-induced bulk refractive index changes. The running buffer was NaH2PO4-Na2HPO4 (10 mM, pH 7.5). Different concentrations (1.25, 2.5, 5, 10, and 20 nM) of xanthan were prepared,and their binding with MiXen-CT was measured by SPR assay. Briefly, the sample solutions were injectedonto the chip surface for 120 s. Following each binding reaction, a further dissociation time of 600 to3,000 s was applied, and as the regeneration buffer, 5 M NaCl solution was injected for 60 s to allow thesignal to return to baseline.

Preparation and chemical analysis of xanthan samples. For xanthan purification, 1 g/liter food-grade xanthan solution in distilled water was precipitated by adding an equal volume of ice-coldabsolute ethanol in the presence of NaCl (100 g/liter). After stirring for 2 h at 4°C, the precipitate wascollected by filtration and washed with 95% ethanol (44). The purified native xanthan was lyophilized forthe following test.

The chemical modification of xanthan was prepared as previously described. Briefly, 5 g/liter purifiedxanthan solution in 5 mM trifluoroacetic acid was heated at 100°C for 1.5 h to remove the pyruvic acetylgroups, which yielded PFX (45). AFX was prepared by saponification treatment of NX with 1 M NaOH at60°C for 1 h to remove the acetyl groups (15). APFX was produced by 5 mM trifluoroacetic acid treatmentfollowed by saponification. To obtain TMFX samples, 10 g/liter purified xanthan solution was incubatedwith 1 mg/ml xanthan lyase, prepared by our laboratory (42), at 40°C for 12 h. All modified xanthanpolymers were dialyzed against demineralized water for 24 h and then lyophilized.

The acetyl content of xanthan was measured according to the method described by Hestrin et al. (46)using acetylcholine as a reference. To determine the pyruvate content of xanthan, 0.01 g xanthan washydrolyzed in 5 ml HCl (1 M) at 100°C for 3 h, after which the free pyruvate was determined as describedby Sloneker et al. (47) using sodium pyruvate as the reference.

Circular dichroism. The ellipticities (millidegrees) of 1 g/liter xanthan or xanthan digests weremonitored at 219 nm using a Jasco J-1500 spectropolarimeter (Jasco Corp., Tokyo, Japan) with a responsetime of 1 s, sensitivity of 100 millidegrees, and bandwidth of 2 nm. The transition profiles of NX, AFX, PFX,APFX, and TMFX were determined in 10 mM NaH2PO4-Na2HPO4 buffer (pH 7.5) at 40°C using a JascoMCB-100 controller. Quartz cuvettes with an optical path of 1 cm were used. The obtained curves werenormalized by the best-fit parameters, and the fractions of disordered conformation (�) were determinedby transition profiles using the following equation:

� � 1 ��t � �U

�F � �U

where �t is ellipticity at a given condition, �U is ellipticity of a completely disordered structure, and �F isellipticity of a completely ordered structure (48). The minimum and maximum ellipticities of each typeof xanthan were determined as described by Kool et al. (8).

Atomic force microscopy imaging of xanthan samples. To allow imaging of individual well-separated xanthan molecules, 5-�l aliquots of undigested or digested xanthan samples (10 mg/liter)were pipetted onto freshly cleaved mica sheets and dried under a gentle flow of dry nitrogen gas.Topographical and error signal mode imaging of the samples was then conducted using a commercialAFM (XE-Bio; Park Systems, South Korea) in noncontact mode with NCHR monolithic silicon cantilevers(Park Systems, South Korea). To provide a disturbance-free environment, an e-Stable mini vibrationisolation table and an acoustic enclosure (Park Systems, South Korea) were used during the measure-ments. The scanning and imaging process was conducted using the AFM controller software XEP(versions 1.7.70.3). The image resolution was set to 256 by 256 pixels for overview images (10 by 10 �m2),and imaging was conducted under ambient conditions with a scan rate of 0.5 Hz.

Raw AFM data were processed and analyzed using the XEI software package (version 1.8.0; ParkSystems, South Korea), while xanthan structural analysis was conducted using the software package XEP(version 1.8.4; Park Systems, South Korea). Using XEI, the raw data were flattened and error lines werecorrected. Furthermore, the height of individual xanthan strands was estimated by XEI. The height of anindividual strand was evaluated by averaging the height estimated from at least five cross sections. Toestimate the mean height, the overall mean of one sample was calculated.

Gel permeation chromatography analysis. The molecular masses of xanthan and its digests weredetermined by gel permeation chromatography (GPC) using an Agilent 1260 infinity system (AgilentTechnologies, Santa Clara, CA, USA). A set of three gel columns (Agilent Technologies, Santa Clara, CA,USA) was used in series with PL aquagel-OH 60, PL aquagel-OH MIXED-M, and PL aquagel-OH 30separation columns (8 �m by 300 mm). The column temperature was set to 40°C, and the samples (20 �l,1 g/liter) were eluted with 0.1 M NaNO3 at a flow rate of 0.8 ml/min (8, 49). The eluate was thenmonitored using refractive index detection. Molecular masses were estimated with the help of pullulanmolecular mass standards (Polymer Laboratories, Palo Alto, CA, USA). The relative molecular massdistribution was calculated by dividing the integrated refractive index (RI) peak area of each fraction bythe total RI peak area measured from 18 to 33 min.

Viscosity measurements. The viscosities of xanthan or xanthan digests were measured against avariety of shear rates using a Discovery hybrid rheometer (TA Instruments), with the following settings:geometry, 40-mm rotating drum; temperature, 40°C; stop head gap, 1,000.0 �m; geometry gap,1,000.0 �m; torque, 5.0 to 1,000.0 �N · m. A total of 1.0 ml of xanthan enzyme products was taken, andthe viscosity was measured against the shear rate for 60 s. All samples were prepared in duplicate.

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Statistical analysis. All analyses were performed in triplicate, and all data were expressed as themeans � standard errors of the means. Analyses were conducted using SPSS 11 (SPSS Inc., Chicago, IL),and a P value of �0.05 was considered significant.

Accession number(s). The amino acid sequences of MiXen and MiGH have been submitted toGenBank under accession numbers ALX66163.1 and WP_067195711, respectively.

SUPPLEMENTAL MATERIALSupplemental material for this article may be found at https://doi.org/10.1128/AEM

.01800-18.SUPPLEMENTAL FILE 1, PDF file, 0.2 MB.

ACKNOWLEDGMENTSFinancial support provided by Natural Sciences Foundation of China (31671796,

31771907, and 31601458), Program for Liaoning Excellent Talents in University(LJQ2015009), Science and Technology Department of Liaoning (201602059), andResearch Initiation Funding for PhDs of Liaoning (201601272) is greatly acknowledged.

REFERENCES1. Lu GT, Ma ZF, Hu JR, Tang DJ, He YQ, Feng JX, Tang JL. 2007. A novel

locus involved in extracellular polysaccharide production and virulenceof Xanthomonas campestris pathovar campestris. Microbiology 153:737–746. https://doi.org/10.1099/mic.0.2006/001388-0.

2. Xiong X, Li M, Xie J, Xue B, Sun T. 2014. Preparation and antioxidantactivity of xanthan oligosaccharides derivatives with similar substitutingdegrees. Food Chem 164:7–11. https://doi.org/10.1016/j.foodchem.2014.05.001.

3. Nankai H, Hashimoto W, Murata K. 2002. Molecular identification offamily 38 �-mannosidase of Bacillus sp. strain GL1, responsible forcomplete depolymerization of xanthan. Appl Environ Microbiol 68:27–31.

4. Hashimoto W, Miki H, Tsuchiya N, Nankai H, Murata K. 2001. Polysac-charide lyase: molecular cloning, sequencing, and overexpression of thexanthan lyase gene of Bacillus sp. strain GL1. Appl Environ Microbiol67:713–720. https://doi.org/10.1128/AEM.67.2.713-720.2001.

5. Hashimoto W, Miki H, Tsuchiya N, Nankai H, Murata K. 1998. Xanthanlyase of Bacillus sp. strain GL1 liberates pyruvylated mannose fromxanthan side chains. Appl Environ Microbiol 64:3765–3768.

6. Thorne L, Tansey L, Pollock TJ. 1987. Clustering of mutations blockingsynthesis of xanthan gum by Xanthomonas campestris. J Bacteriol 169:3593–3600. https://doi.org/10.1128/jb.169.8.3593-3600.1987.

7. Cadmus MC, Jackson LK, Burton KA, Plattner RD, Slodki ME. 1982.Biodegradation of xanthan gum by Bacillus sp. Appl Environ Microbiol44:5–11.

8. Kool MM, Schols HA, Delahaije RJBM, Sworn G, Wierenga PA, Gruppen H.2013. The influence of the primary and secondary xanthan structure onthe enzymatic hydrolysis of the xanthan backbone. Carbohydr Polym97:368 –375. https://doi.org/10.1016/j.carbpol.2013.05.045.

9. Cheetham NWH, Mashimba ENM. 1991. Characterisation of some enzy-mic hydrolysis products of xanthan. Carbohydr Polym 15:195–206.https://doi.org/10.1016/0144-8617(91)90032-8.

10. Christensen BE, Smidsrød O. 1996. Dependence of the content of un-substituted (cellulosic) regions in prehydrolysed xanthans on the rate ofhydrolysis by Trichoderma reesei endoglucanase. Int J Biol Macromol18:93–99. https://doi.org/10.1016/0141-8130(95)01063-7.

11. Rinaudo M, Milas M. 1980. Enzymic hydrolysis of the bacterial polysac-charide xanthan by cellulase. Int J Biol Macromol 2:45– 48. https://doi.org/10.1016/0141-8130(80)90009-4.

12. Matsuda Y, Biyajima Y, Sato T. 2009. Thermal denaturation, renatur-ation, and aggregation of a double-helical polysaccharide xanthan inaqueous solution. Polym J 41:526 –532. https://doi.org/10.1295/polymj.PJ2008300.

13. Bezemer L, Ubbink JB, Kooker JAD, Kuil ME, Leyte JC. 1993. On theconformational transitions of native xanthan. Macromolecules 26:6436 – 6446. https://doi.org/10.1021/ma00076a021.

14. Rinaudo M. 2004. Role of substituents on the properties of some poly-saccharides. Biomacromolecules 5:1155–1165. https://doi.org/10.1021/bm030077q.

15. Shatwell KP, Sutherland IW, Dea ICM, Ross-Murphy SB. 1990. The influ-ence of acetyl and pyruvate substituents on the helix-coil transition

behaviour of xanthan. Carbohydr Res 206:87–103. https://doi.org/10.1016/0008-6215(90)84009-J.

16. Qian F, An L, Wang M, Li C, Li X. 2007. Isolation and characterizationof a xanthan-degrading Microbacterium sp. strain XT11 from gardensoil. J Appl Microbiol 102:1362–1371. https://doi.org/10.1111/j.1365-2672.2006.03215.x.

17. Fan Y, Li L, Yang S, Ming Y, Guo X, Hou Y, Chen X, Li X. 2016. Completegenome sequence of a xanthan-degrading Microbacterium sp. strainXT11 with the potential for xantho-oligosaccharides production. J Bio-technol 222:19 –20. https://doi.org/10.1016/j.jbiotec.2016.02.005.

18. Okano H, Kanaya E, Ozaki M, Angkawidjaja C, Kanaya S. 2015. Structure,activity, and stability of metagenome-derived glycoside hydrolase family9 endoglucanase with an N-terminal Ig-like domain. Protein Sci 24:408 – 419. https://doi.org/10.1002/pro.2632.

19. Li B, Guo J, Chen W, Chen X, Chen L, Liu Z, Li X. 2009. Endoxanthanase,a novel �-D-glucanase hydrolyzing backbone linkage of intact xanthanfrom newly isolated Microbacterium sp. XT11. Appl Biochem Biotechnol159:24 –32. https://doi.org/10.1007/s12010-008-8439-1.

20. Teckentrup J, Al-Hammood O, Steffens T, Bednarz H, Walhorn V, NiehausK, Anselmetti D. 2017. Comparative analysis of different xanthan sam-ples by atomic force microscopy. J Biotechnol 257:2– 8. https://doi.org/10.1016/j.jbiotec.2016.11.032.

21. Qi M, Jun H-S, Forsberg CW. 2007. Characterization and synergisticinteractions of fibrobacter succinogenes glycoside hydrolases. Appl En-viron Microbiol 73:6098 – 6105. https://doi.org/10.1128/AEM.01037-07.

22. Arai T, Araki R, Tanaka A, Karita S, Kimura T, Sakka K, Ohmiya K. 2003.Characterization of a cellulase containing a family 30 carbohydrate-binding module (CBM) derived from Clostridium thermocellum CelJ:importance of the CBM to cellulose hydrolysis. J Biotechnol 185:504 –512. https://doi.org/10.1128/JB.185.2.504-512.2003.

23. Eckert K, Zielinski F, Lo LL, Schneider E. 2002. Gene cloning, sequencing,and characterization of a family 9 endoglucanase (CelA) with an unusualpattern of activity from the thermoacidophile Alicyclobacillus acidocal-darius ATCC27009. Appl Microbiol Biotechnol 60:428 – 436. https://doi.org/10.1007/s00253-002-1131-4.

24. Hirano N, Hasegawa H, Nihei S, Haruki M. 2013. Cell-free protein syn-thesis and substrate specificity of full-length endoglucanase CelJ (Cel9D-Cel44A), the largest multi-enzyme subunit of the Clostridium thermo-cellum cellulosome. FEMS Microbiol Lett 344:25–30. https://doi.org/10.1111/1574-6968.12149.

25. Ravachol J, Borne R, Tardif C, Philip PD, Fierobe HP. 2014. Characteriza-tion of all family-9 glycoside hydrolases synthesized by the cellulosome-producing bacterium Clostridium cellulolyticum. J Biol Chem 289:7335–7348. https://doi.org/10.1074/jbc.M113.545046.

26. Gulrez SKH, Al-Assaf S, Fang Y, Phillips GO, Gunning AP. 2012. Revisitingthe conformation of xanthan and the effect of industrially relevanttreatments. Carbohydr Polym 90:1235–1243. https://doi.org/10.1016/j.carbpol.2012.06.055.

27. Araki Y, Karita S, Tanaka A, Kondo M, Goto M. 2009. Characterization offamily 17 and family 28 carbohydrate-binding modules from Clostridium

Insights into Xanthan Hydrolysis by Xanthanase MiXen Applied and Environmental Microbiology

January 2019 Volume 85 Issue 2 e01800-18 aem.asm.org 15

on August 28, 2020 by guest

http://aem.asm

.org/D

ownloaded from

josui Cel5A. Biosci Biotechnol Biochem 73:1028 –1032. https://doi.org/10.1271/bbb.80802.

28. Tsukimoto K, Takada R, Araki Y, Suzuki K, Karita S, Wakagi T, Shoun H,Watanabe T, Fushinobu S. 2010. Recognition of cellooligosaccharides bya family 28 carbohydrate-binding module. FEBS Lett 584:1205–1211.https://doi.org/10.1016/j.febslet.2010.02.027.

29. Reyes-Ortiz V, Heins RA, Cheng G, Kim EY, Vernon BC, Elandt RB, AdamsPD, Sale KL, Hadi MZ, Simmons BA, Kent MS, Tullman-Ercek D. 2013.Addition of a carbohydrate-binding module enhances cellulase pene-tration into cellulose substrates. Biotechnol Biofuels 6:93–106. https://doi.org/10.1186/1754-6834-6-93.

30. Nakamura A, Tasaki T, Ishiwata D, Yamamoto M, Okuni Y, Visootsat A,Maximilien M, Noji H, Uchiyama T, Samejima M, Igarashi K, Iino R. 2016.Single-molecule imaging analysis of binding, processive movement, anddissociation of cellobiohydrolase Trichoderma reesei Cel6A and its do-mains on crystalline cellulose. J Biol Chem 291:22404 –22413. https://doi.org/10.1074/jbc.M116.752048.

31. Ståhlberg J, Johansson G, Pettersson G. 1991. A new model for enzy-matic hydrolysis of cellulose based on the two-domain structure ofcellobiohydrolase I. Nat Biotechnol 9:286 –290. https://doi.org/10.1038/nbt0391-286.

32. Várnai A, Mäkelä MR, Djajadi DT, Rahikainen J, Hatakka A, Viikari L.2014. Carbohydrate-binding modules of fungal cellulases: occurrencein nature, function, and relevance in industrial biomass conversion.Adv Appl Microbiol 88:103–165. https://doi.org/10.1016/B978-0-12-800260-5.00004-8.

33. Varnai A, Viikari L, Siika-Aho M. 2013. Carbohydrate-binding modules(CBMs) revisited: reduced amount of water; counterbalances the needfor CBMs. Biotechnol Biofuels 6:30 – 42. https://doi.org/10.1186/1754-6834-6-30.

34. Arendt O, Kulicke WM. 1998. Determination of the viscoelastic propertiesof a homologous series of the fermentation polymer xanthan gum.Angew Makromol Chem 259:61– 67. https://doi.org/10.1002/(SICI)1522-9505(19981001)259:1�61::AID-APMC61�3.0.CO;2-V.

35. Smith IH, Symes KC, Lawson CJ, Morris ER. 1981. Influence of thepyruvate content of xanthan on macromolecular association in solu-tion. Int J Biol Macromol 3:129 –134. https://doi.org/10.1016/0141-8130(81)90078-7.

36. Tako M, Nakamura S. 1984. Rheological properties of deacetylated xan-than in aqueous media. Agric Biol Chem 48:2987–2993. https://doi.org/10.1080/00021369.1984.10866637.

37. Moorhouse R, Walkinshaw MD, Arnott S. 1977. Xanthan gum–molecular

conformation and interactions. Extracell Microb Polysaccharides 45:90 –102. https://doi.org/10.1021/bk-1977-0045.

38. Yin Y, Mao X, Yang J, Chen X, Mao F, Xu Y. 2012. dbCAN: a web resourcefor automated carbohydrate-active enzyme annotation. Nucleic AcidsRes 40:445– 451.

39. Quevillon E, Silventoinen V, Pillai S, Harte N, Mulder N, Apweiler R, LopezR. 2005. InterProScan: protein domains identifier. Nucleic Acids Res33:116 –120.

40. Roy A, Kucukural A, Zhang Y. 2010. I-TASSER: a unified platform forautomated protein structure and function prediction. Nat Protoc5:725–738. https://doi.org/10.1038/nprot.2010.5.

41. Crowe J, Dobeli H, Gentz R, Hochuli E, Stiiber D, Henco K. 1994. 6xHis-Ni-NTA chromatography as a superior technique in recombinant proteinexpression/purification. Methods Mol Biol 31:371–387.

42. Yang F, Yang L, Guo X, Wang X, Li L, Liu Z, Wang W, Li X. 2014.Production and purification of a novel xanthan lyase from a xanthan-degrading Microbacterium sp. strain XT11. Sci World J 2014:1. ID368434.https://doi.org/10.1155/2014/368434.

43. Mcfeeters RF. 1980. A manual method for reducing sugar determinationswith 2,2’-bicinchoninate reagent. Anal Biochem 103:302–306. https://doi.org/10.1016/0003-2697(80)90614-4.

44. Callet F, Milas M, Rinaudo M. 1987. Influence of acetyl and pyruvate con-tents on rheological properties of xanthan in dilute solution. Int J BiolMacromol 9:291–293. https://doi.org/10.1016/0141-8130(87)90068-7.

45. Bradshaw IJ, Nisbet BA, Kerr MH, Sutherland IW. 1983. Modified xanthan-its preparation and viscosity. Carbohydr Polym 3:23–38. https://doi.org/10.1016/0144-8617(83)90010-3.

46. Hestrin S. 1949. The reaction of acetylcholine and other carboxylic acidderivatives with hydroxylamine, and its analytical application. J BiolChem 180:249 –261.

47. Sloneker JH, Orentas DG. 1962. Pyruvic acid, a unique component of anexocellular bacterial polysaccharide. Nature 194:478 – 479. https://doi.org/10.1038/194478a0.

48. Greenfield NJ. 2007. Using circular dichroism collected as a function oftemperature to determine the thermodynamics of protein unfoldingand binding interactions. Nat Protoc 1:2527–2535. https://doi.org/10.1038/nprot.2006.204.

49. Milas M, Rinaudo M, Tinland B. 1986. Comparative depolymerization ofxanthan gum by ultrasonic and enzymic treatments. Rheological andstructural properties. Carbohydr Polym 6:95–107. https://doi.org/10.1016/0144-8617(86)90037-8.

Yang et al. Applied and Environmental Microbiology

January 2019 Volume 85 Issue 2 e01800-18 aem.asm.org 16

on August 28, 2020 by guest

http://aem.asm

.org/D

ownloaded from