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Originally published In Press as doi:10.1074/jbc.M700224200 on March 21, 2007

J. Biol. Chem., Vol. 282, Issue 26, 19177-19189, June 29, 2007
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Characterization and Three-dimensional Structures of Two Distinct Bacterial Xyloglucanases from Families GH5 and GH12*

Tracey M. Gloster{ddagger}1, Farid M. Ibatullin§1, Katherine Macauley{ddagger}1, Jens M. Eklöf§, Shirley Roberts{ddagger}, Johan P. Turkenburg{ddagger}, Mads E. Bjørnvad, Per Linå Jørgensen, Steffen Danielsen, Katja S. Johansen, Torben V. Borchert, Keith S. Wilson{ddagger}, Harry Brumer§2, and Gideon J. Davies{ddagger}3

From the {ddagger}York Structural Biology Laboratory, Department of Chemistry, University of York, York YO10 5YW, United Kingdom, §School of Biotechnology, Royal Institute of Technology (KTH), AlbaNova University Centre, 106 91 Stockholm, Sweden, and Novozymes A/S, Brudelysvej 26, 1U1.23, DK-2880 Bagsvaerd, Denmark

Received for publication, January 9, 2007 , and in revised form, March 9, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The plant cell wall is a complex material in which the cellulose microfibrils are embedded within a mesh of other polysaccharides, some of which are loosely termed "hemicellulose." One such hemicellulose is xyloglucan, which displays a beta-1,4-linked D-glucose backbone substituted with xylose, galactose, and occasionally fucose moieties. Both xyloglucan and the enzymes responsible for its modification and degradation are finding increasing prominence, reflecting both the drive for enzymatic biomass conversion, their role in detergent applications, and the utility of modified xyloglucans for cellulose fiber modification. Here we present the enzymatic characterization and three-dimensional structures in ligand-free and xyloglucan-oligosaccharide complexed forms of two distinct xyloglucanases from glycoside hydrolase families GH5 and GH12. The enzymes, Paenibacillus pabuli XG5 and Bacillus licheniformis XG12, both display open active center grooves grafted upon their respective (beta/{alpha})8 and beta-jelly roll folds, in which the side chain decorations of xyloglucan may be accommodated. For the beta-jelly roll enzyme topology of GH12, binding of xylosyl and pendant galactosyl moieties is tolerated, but the enzyme is similarly competent in the degradation of unbranched glucans. In the case of the (beta/{alpha})8 GH5 enzyme, kinetically productive interactions are made with both xylose and galactose substituents, as reflected in both a high specific activity on xyloglucan and the kinetics of a series of aryl glycosides. The differential strategies for the accommodation of the side chains of xyloglucan presumably facilitate the action of these microbial hydrolases in milieus where diverse and differently substituted substrates may be encountered.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Xyloglucans comprise a family of plant polysaccharides united by a common beta(1->4) glucan backbone regularly decorated at C-6 with {alpha}-linked xylopyranosyl residues (1). Numerous studies have indicated that most xyloglucans are based upon the Glc4 oligosaccharide repeats XXXG or XXGG, where G and X denote unsubstituted D-Glcp and {alpha}-D-Xylp(1->6)-D-Glcp units, respectively (2). More rarely, some species have been observed to produce xyloglucans with both Glc4 and Glc5 backbone repeats, such as XXXXG (3) and XXGGG (4). The xylose branches may in turn be substituted with combinations of galactopyranose, fucopyranose, arabinofuranose, and O-acetyl residues, depending upon the plant species and tissue localization (reviewed in Refs. 2, 5, and 6). A concise, linear notation based on single-letter abbreviations of commonly observed microstructures is widely used to simplify the description of xyloglucans and xylogluco-oligosaccharides (7) as follows: G, D-Glcp; X, {alpha}-D-Xylp(1->6)-D-Glcp;L, beta-D-Galp-(1->2)-{alpha}-D-Xylp(1->6)-D-Glcp; S, {alpha}-L-Araf(1->2)-{alpha}-D-Xylp(1->6)-D-Glcp;F, {alpha}-L-Fucp(1->2)-beta-D-Galp-(1->2)-{alpha}-D-Xylp(1->6)-D-Glcp. The general structure of xyloglucans based on XXXG repeat units, such as the widely studied xyloglucan from tamarind (Tamarindus indica) seed kernel, is shown in Fig. 1.

In plants, xyloglucans function both as seed storage carbohydrates (8) and as essential modulators of the mechanical properties of the primary cell wall (see Refs. 9-14 and references therein). In the latter context, xyloglucans are intimately associated with cellulose through surface adsorption and direct entrapment within the paracrystalline structure (15). Primary cell wall xyloglucans are widely distributed among land plants (16), thus suggesting that the specific cellulose-xyloglucan interaction may have conferred a particular structural advantage in the colonization of drier habitats (17, 18).

There is strong and evolving interest in xyloglucans and also in the enzymes responsible for their modification and degradation. Such interest stems not only from the role of these polysaccharides and their catalysts in plant cell wall morphogenesis (9, 10, 13), but also from biotechnological applications as diverse as fruit juice clarification (19, 20), textile processing (21, 22), cellulose surface modification (23-27), pharmaceutical delivery (28-30), production of food thickening agents (30, 31), as well as the production of xylogluco-oligosaccharides for cell wall analysis (4, 5, 32), plant growth modulation (33, 34), surfactant synthesis (35), and enzyme kinetic studies (36). Furthermore, the goal of biofuel production from plant biomass, which strives to substantially reduce fossil fuel usage, has caused a great resurgence of interest in plant cell wall degrading enzymes (37). However, plant biomass remains extremely difficult to exploit, primarily because its components are extremely resistant to degradation; plant cell wall polysaccharides are often present as insoluble, cross-linked structures. Furthermore, the chemistry of the glycosidic bond itself makes its hydrolysis one of the most challenging reactions in nature, with Wolfenden showing that, in the absence of biological catalysts for its degradation, cellulose has a half-life in excess of 4 million years (38).

The biocatalysts responsible for the hydrolysis of the backbone of xyloglucan are xyloglucan endo-beta-1,4-glucanases or "xyloglucanases" (EC 3.2.1.151 [EC] ). This enzyme commission number reflects many different enzyme sequences, structures, and hydrolytic mechanisms with either inversion or retention of the configuration of the anomeric carbon. In the sequence-based CAZy (carbohydrate active enzymes) classification (39) (recently reviewed in Ref. 40), enzymes defined as xyloglucanases are found in retaining families GH5, GH12, and GH16 and inverting families GH44 and GH74. To date, xyloglucanase structures have only been reported for the GH74 family enzymes (41, 42), although the coordinates for a Clostridial xyloglucanase Cel44A are deposited but currently unavailable (Protein Data Bank code 2D8G).4 The structure of the poplar xyloglucan endotransglycosylase (EC 2.4.1.207 [EC] ) from family GH16, which strongly favors transglycosylation of xyloglucan over hydrolysis, has also been determined (43).

Here we report the characterization, both on polymeric substrates and defined aryl xyloglucan oligosaccharides (Fig. 1), of two structurally distinct xyloglucan hydrolases from families GH5 and GH12: P. pabuli XG5 (hereafter PpXG5) and B. licheniformis XG12 (hereafter BlXG12). The single crystal x-ray structures of both enzymes have been determined, at resolutions from 1.95 to 1.40 Å, in both an unliganded form and in complex with xyloglucan oligosaccharides based upon a cellotetraose backbone. These three-dimensional complexes interpreted in light of the kinetics of the two enzyme classes on xyloglucan-derived substrates provide an unique insight into the different ways xyloglucan side chains are accommodated and/or harnessed for catalysis.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cloning and Expression of PpXG5
The 40-kDa xyloglucanase produced by the P. pabuli strain (DSM 13330) was cloned by standard methods. Briefly, purified genomic P. pabuli DNA was partially digested by Sau3A and cloned into an Escherichia coli-based lambdaZAPexpress vector (Stratagene, La Jolla, CA). Ligated DNA was packaged in phages using the GigaPackIII gold kit (Stratagene). Eventually, plaque-forming phages were screened on agar plates containing AZCL-xyloglucan (Megazyme International Ireland Ltd.), and positive clones were seen by the formation of blue halos. The gene encoding PpXG5 was DNA-sequenced, and PCR primers were designed for amplification of the gene from P. pabuli genomic DNA. The PpXG5 gene was cloned into a B. subtilis expression vector and expressed as a secreted form from the amyl promoter. PpXG5 was recovered from the broth through a combination of chemical and physical separation steps. The supernatant was applied to a pre-equilibrated S-Sepharose column at pH 5 in 20 mM sodium acetate buffer. PpXG5 was eluted with a gradient of 1 M NaCl in 20 mM sodium acetate, and the appropriate fractions were pooled. PpXG5 was further purified by gel filtration on an S200 column in 0.1 M sodium acetate buffer, pH 6.

Cloning and Expression of BlXG12
The 26-kDa xyloglucanase produced by the B. licheniformis strain (ATCC14580) was cloned from B. licheniformis genomic DNA, and the gene was expressed in B. subtilis by essentially the same protocol as described above for PpXG5. Protein purification also followed a similar strategy. Enzymatic variants of BlXG12 were constructed using the megapriming method and purified as for the wild type enzyme.

Kinetic Characterization of PpXG5 and BlXG12
Substrate Specificity—Substrate specificity was determined using the Somogyi-Nelson reducing sugar assay (44, 45) (total assay volume 100 µl, 700 µl of reagents added) versus a standard curve for glucose (1.0-15 µg). The following polysaccharides were used: konjac glucomannan, carboxymethyl cellulose, Alcaligenes faecalis curdlan, wheat arabinoxylan, carob galactomannan (low viscosity), Icelandic moss lichenan, tamarind xyloglucan, and barley beta-glucan (all from Megazyme); Avicel, hydroxyethyl cellulose, and apple pectin (70-75% esterification) (Fluka); and birchwood xylan and citrus pectic acid (Sigma). Soluble polysaccharides were tested in triplicate at 0.5 g/liter by incubation with each enzyme (0.23 mg/liter PpXG5 or 5.9 mg/liter BlXG12) at 55 °C in 50 mM NaOAc buffer, pH 5.5, for 15 min (initial rate conditions). Reducing sugars were quantified at 520 nm with a Cary 50 UV-visible spectrophotometer (Varian).

pH Rate Profiles—The pH rate profiles of PpXG5 and BlXG12 were determined in triplicate using the method described by Nelson and Somogyi with tamarind xyloglucan (1 g/liter) as the substrate. The following 50 mM buffer systems were used: sodium acetate, pH 4.0-5.5, and sodium phosphate, pH 5.75-8.

Time-dependent Depolymerization of Xyloglucan—Samples (600-µl total volume) containing tamarind xyloglucan (1 g/liter) and PpXG5 (56 µg/liter) or BlXG12 (840 µg/liter) in 50 mM sodium acetate buffer, pH 5.5, were aliquoted and removed after incubation for 7, 15, 30, 45, and 60 min at 55 °C or overnight incubation at 37 °C. Samples were freeze-dried and dissolved in dimethyl sulfoxide before size exclusion chromatography on an HPLC5 system composed of a Gynkotec 480 pump, a Gynkotec Gina 50 autosampler, two Tosoh gel columns, G5000HHR and G3000HHR (both 7.8 x 300 mm), connected in series, and an evaporative light-scattering detector (PL-ELS 1000; Polymer Laboratories). HPLC grade Me2SO was used as the eluent at a flow rate of 1.0 ml/min, and the column temperature was maintained at 60 °C (27).


Figure 1
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  		FIGURE 1.
General structure of XXXG-based xyloglucans and chromogenic xyloglucanase substrates. A, endoglucanases, including PpXG5 and BlXG12, typically cleave the glycosidic bond of the unbranched Glc residue (arrow) to yield xylogluco-oligosaccharides. Common structures in seed xyloglucans, such as that from T. indica, include XXXG (x = 0, y = 0, R = H), XLXG (x = 1, y = 0, R = H), XXLG (x = 0, y = 1, R = H), and XLLG (x = 1, y = 1, R = H). XXFG (x = 0, y = 1, R = {alpha}-1,2-L-Fuc) is prevalent in xyloglucans from dicot primary cell walls. B, the chromogenic aryl beta-glycosides of xylogluco-oligosaccharides employed for detailed kinetic analysis of PpXG5 and BlXG12 were XXXG-CNP (x = 0, y = 0, R'= Cl, R''= NO2), XLLG-CNP (x = 1, y = 1, R'= Cl, R''= NO2), and XXXG-pMP (x = 0, y = 0, R'= H, R''= OCH3).

 
Limit Digest Analysis—Extended enzymatic hydrolysis of xyloglucan (0.5 g/liter) in 50 mM NaOAc buffer, pH 5.5, was performed at 37 °C overnight with PpXG5 (280 µg/liter) or BlXG12 (58,000 µg/liter). Samples were analyzed with a Dionex ICS-3000 high performance anion exchange chromatography system with pulsed amperometric detection (HPAEC-PAD) and a Dionex PA-100 column using a gradient modified from that previously described (41). Conditions were as follows: Solvent A, 1.0 M NaOH; solvent B, 1.0 M NaOAc; Solvent C, ultrapure water; flow rate, 0.8 ml/min. The gradient program was as follows: 0-3 min, 100 mM NaOH, 40 mM NaOAc; 3-18 min, linear gradient from 40 to 300 mM NaOAc; 18-19 min, gradient up to 500 mM NaOH and 500 mM NaOAc and then initial conditions for 4 min.

Hydrolysis of Aryl (Xylo)glucooligosaccharides—The 2-chloro-4-nitrophenyl (CNP) and 4-methoxylphenyl (pMP) beta-glycosides of GGGG (cellotetraose), XXXG, and XLLG (Fig. 1) were synthesized from the corresponding per-O-acetyl derivatives (35) by a sequence of anomeric {alpha}-bromination, phase transfer glycosylation with the corresponding phenolate, and deprotection under Zemplén conditions; full experimental details will be published elsewhere.6 All XGO aryl glycosides were purified on C18-silica reversed-phase columns (RP18; Supelco) before use and exhibited satisfactory NMR and mass spectra.

The enzymatic hydrolyses of GGGG-CNP, XXXG-CNP, and XLLG-CNP were followed by continuous assays measuring the release of 2-chloro-4-nitrophenolate at 405 nm (measured {epsilon} 9724 M-1 cm-1, 5 mM NaOAc buffer, pH 5.5) using a Cary 300 Bio UV-visible spectrophotometer (Varian). A total assay volume of 100 µl was used in 1-cm path length quartz cells equilibrated and maintained at 30 ± 0.1 °C in a Peltier-controlled cell block. Initial rates were determined from the slope of the linear region of the reaction time course corresponding to <10% conversion. Assays of PpXG5 and BlXG12 employed total enzyme (protein) concentrations of 1.4 µg/ml (0.035 µM) and 46.8 µg/ml (1.79 µM), respectively. Kinetic constants were obtained from plots of vo/[E]t versus [S] by nonlinear curve fitting using MicrocalTM Origin® version 6.0. In the absence of an active site titrant, [E]t was assumed to be equivalent to the total protein concentration (i.e. 100% active protein).

The rates of enzyme-catalyzed hydrolysis of XXXG-pMP were determined by incubation in 5.0 mM NaOAc buffer, pH 5.5, with the temperature maintained in a thermostated block at 30 ± 0.1 °C (100-µl total assay volume). The incubation time for PpXG5 was 60 min, and the concentration of the enzyme in the assay was 1.4 µg/ml (0.035 µM). For BlXG12, the incubation time was 60 min, and the enzyme concentration was 117 µg ml-1 (4.49 µM). The reactions were stopped by the addition of 0.2 M sodium carbonate (100 µl). The concentration of released 4-methoxyphenolate was determined by measuring the absorbance at 305 nm ({epsilon} 2656 M-1 cm-1) using a Cary 50 Bio UV-visible spectrophotometer (Varian) and subtraction of background values from XXXG-pMP.

Crystallization, Data Collection, and Structure Solution of PpXG5
PpXG5, at ~15 mg/ml, was crystallized from 20-25% (w/v) polyethylene glycol 8000, 0.2 M CaCl2 and 0.1 M Tris-HCl, pH 8.5. Single crystals were cryoprotected with mother liquor with the addition of 25% (v/v) ethylene glycol. Data were collected to 1.40 Å on ESRF beamline ID14-3 and processed with DENZO/SCALEPACK (46). The structure was solved by molecular replacement using the Clostridium cellulolyticum Cel5A (Cel-CCA) as the search model (Protein Data Bank code 1EDG) (47) with the CCP4 (48) version of the program AMORE (49), using the default parameters. An initial model was built automatically with the CCP4 installation of ARP-wARP, and the structure was refined using REFMAC (50) with manual corrections using QUANTA (Accelrys, San Diego, CA) and COOT (51). Data and structure statistics are given in Table 1.


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  		TABLE 1
Data collection and refinement statistics

 
Structure of a Xyloglucan Oligosaccharide Complex of PpXG5
PpXG5, in 25 mM acetate buffer, pH 6, was crystallized in the presence of ~10 mM mixed xyloglucan oligosaccharides (2:1: 3:3 mixture of XXXG/XLXG/XXLG/XLLG as described in Ref. 41) from 25% polyethylene glycol 3350, 0.2 M MgCl2, 0.1 M Tris/HCl, pH 8.5, and 0.01 M Tris. Crystals were cryoprotected using the crystal growth conditions with the addition of 5% ethylene glycol prior to flash-freezing in liquid nitrogen.

Data were collected at the ESRF on beamline ID14-1 to 1.95 Å resolution from a single crystal cooled to 100 K. Data were integrated and processed with DENZO and scaled and merged with SCALEPACK in HKL2000 (46). All subsequent computing was done using the CCP4 suite of programs (48). The structure of the PpXG5 complex was solved by molecular replacement with the native structure of PpXG5 as the search model, using the CCP4 version of AMORE (49) with data between 15 and 3 Å and an outer radius of Patterson integration of 25 Å. The structure was subsequently refined using REFMAC (50) interspersed with manual corrections and the addition of waters in COOT (51).

Crystallization, Data Collection, and Structure Solution of BlXG12
Initial crystals of the native form of BlXG12 were obtained in a 50:50 merohedrally twinned crystal form (apparent space group P4n a = b = 124 Å, c = 50 Å; data to 2.2 Å, Rmerge = 10% at edge). Subsequently, a E155Q variant of BlXG12 was used to obtain a nontwinned crystal form. BlXG12(E155Q) was crystallized from 10% (w/v) polyethylene glycol 4000, 0.05 M MgCl2, and 0.1 M BisTris, pH 6.5. Native data were collected in the home laboratory (YSBL, York, UK) using CuK{alpha} radiation and a MARResearch Image Plate system. Data were processed with DENZO/SCALEPACK (46), and the structure was solved with the Streptomyces lividans Cel12 (52, 53) as the search model. ARP-wARP was unable to build this model automatically, so partial manual rebuilding and refinement with REFMAC (50) were required to produce a starting model within the convergence radius of ARP-wARP. Following completion of a partial model, refinement continued with REFMAC with manual corrections using QUANTA (Accelrys) and COOT (51).

Structure of a Ligand Complex of BlXG12 (E155A)
BlXG12 E155A, in 25 mM acetate buffer and in the presence of ~10 mM mixed xyloglucan oligosaccharides, was crystallized from 1.6 M ammonium sulfate, 10% dioxane, and 0.1 M 2-(N-morpholino)-ethanesulfonic acid, pH 6.5. The mother liquor with the addition of 23% glycerol was used to cryoprotect the crystal prior to flash-freezing in liquid nitrogen.

Data were collected at the ESRF on beamline ID14-1 from a single crystal cooled to 100 K to 1.40 Å resolution. The structure was solved using AMORE (49) with the unliganded BlXG12 E155Q mutant as the search model, and was refined as described previously for the PpXG5 complex.


Figure 2
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  		FIGURE 2.
High performance size exclusion (A) and anion exchange (B) chromatographic analysis of the limit digestion of tamarind xyloglucan by PpXG5.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
To discover novel xyloglucan-degrading enzymes, fragmented genomic DNA from both B. licheniformis and P. pabuli were cloned into E. coli and bacteriophage {lambda} expression vectors, and the subsequent libraries were screened for the expression of xyloglucan active enzymes by plating onto AZCL-xyloglucan-agarose. Using this strategy, BlXG12 and PpXG5 were discovered with initial data on dyed xyloglucans, demonstrating that these enzymes could be classified as "xyloglucanases."

Despite family GH5 having over 900 members, the PpXG5 enzyme is almost unique, with only two sequences from Paenibacillus sp. KM21 and Bacillus sp. BP-23 having >45% sequence identity. The former has been shown to be an obligate xyloglucanase (54), whereas the latter finds use in straw processing applications (55) but has not, to our knowledge, been tested on xyloglucan substrates. Family GH12 has ~110 members. BlXG12 shows high (>50%) sequence identity with only one other enzyme, that from Pectobacterium carotovorum (Erwinia carotovora) with ~67% identity, which has been described as an endoglucanase, but again activity on xyloglucan was not reported (56, 57). The next most similar sequences are those from Streptomyces avermitilis (41% identity) and EG12 from Rhodothermus marinus (30% identity), whose three-dimensional structure has previously been reported (58).

Kinetic Analysis of PpXG5—Relative hydrolysis rates indicate that PpXG5 is an exclusive xyloglucanase, with no detectable activity on a range of other glucan, xylan, mannan, or pectic polysaccharides (Table 2), as observed recently for a close homolog (54). The dependence of the enzymatic hydrolysis rate of xyloglucan on pH was classically bell-shaped (data not shown), with apparent kinetic pKa values of two ionizable groups of 4.5 ± 0.2 and 8.0 ± 0.2. Size exclusion chromatography and HPAEC-PAD demonstrated that PpXG5 hydrolyzes tamarind xyloglucan endolytically to produce a mixture of the Glc4-based oligosaccharides XXXG, XLXG, XXLG, and XLLG (Fig. 2).


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  		TABLE 2
Kinetics of PpXG5 and BlXG12 on polysaccharide substrates

ND, activity not detected. Neither enzyme showed activity on hydroxyethyl cellulose, carob galactomannan, Icelandic moss lichenan, wheat or birchwood xylans, apple pectin, or citrus pectic acid. Errors shown are the S.D. values derived from triplicate determinations.

 
PpXG5 was also active on synthetic CNP (59), beta-glycosides of GGGG, XXXG, and XLLG, and the pMP beta-glycoside of XXXG (Fig. 1B). In the case of GGGG-CNP, XXXG-CNP, and XXXG-pMP, PpXG5 exhibited classical saturation kinetics, and the data (Fig. 3) were readily fit by the standard Michaelis-Menten equation. The rate of the PpXG5-catalyzed hydrolysis of XLLG-CNP, however, showed a more complex dependence on substrate concentration (Fig. 4), which indicated that the glycosylenzyme intermediate, Egly, was capable of binding a second molecule of substrate at high [S], giving rise to substrate inhibition. This was fit appropriately to yield the substrate inhibition constant Kis, in addition to kcat and Km (Table 3), using V = Vmax [S]/Km + [S] + ([S]2/Ki).


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  		TABLE 3
Kinetics of PpXG5 and BIXG12 on aryl (xylo)gluco-oligosaccharide beta-glycosides

Kinetic constants were obtained by nonlinear least squares curve fitting.

 


Figure 3
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  		FIGURE 3.
Initial rate kinetics of the PpXG5-catalyzed hydrolysis of XXXG-CNP. A, low substrate concentration regime; B, full substrate concentration range. Lines represent nonlinear least squares fits of the standard Michaelis-Menten equation to the data.

 


Figure 4
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  		FIGURE 4.
Initial rate kinetics of the PpXG5-catalyzed hydrolysis of XLLG-CNP. A, low substrate concentration regime; B, full substrate concentration range. Lines represent nonlinear least squares fits incorporating substrate inhibition.

 
The degree of substrate inhibition is quite low for XLLG-CNP and is only manifested at substrate concentrations well above the apparent Km value (Fig. 4, Table 3). GH5 enzymes are retaining, with catalysis occurring via the formation and subsequent breakdown of a covalent glycosyl-enzyme intermediate. With a good leaving group, such as 2-chloro-4-nitrophenol (pKa = 5.45) (60), the low observed Km value is indicative of rate-determining breakdown of the covalent glycosyl-enzyme (k2 >> k3). Indeed, this is supported by the Km value of XXXG-pMP (4-methoxypenolate pKa = 10.21) (61), which is 7-fold higher than for XXXG-CNP. Similar leaving group effects on Km values and the rate-determining step have been previously observed in glycanases, for example with the endo-xylanase from Cellulomonas fimi (60). Kinetic and product analysis data are consistent with the accumulation of an XLLG-PpXG5 glycosyl-enzyme intermediate that binds a second molecule of substrate to form a dead end complex; no evidence for transglycosylation leading to formation of XLLGXLLG-CNP or higher oligomers was observed by HPAEC-PAD.


Figure 5
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  		FIGURE 5.
A, three-dimensional protein schematic representation of PpXG5 in complex with XXLG. The protein chain is shown color-ramped from the N terminus (blue) to C terminus (red) with the ligand and the catalytic acid/base and nucleophile in a ball-and-stick representation. B, observed electron density, interpreted as a single molecule of XXLG bound to the -4 to -1 subsites of the enzyme. Both images, drawn with MOLSCRIPT (68)/BOBSCRIPT (69), are in divergent (wall-eyed) stereo representation.

 
The values of the macroscopic kinetic constants obtained for the action of PpXG5 on XXXG-CNP and XLLG-CNP are summarized in Table 3. Although both substrates have Km values in the micromolar range, the Km value for XLLG-CNP is 2-fold lower. The ratio of kcat/Km indicates that PpXG5 is more selective for this substrate than XXXG-CNP by a factor of ~3. The presence of one or both additional galactose residues thus enhances catalysis by the enzyme, lowering the activation barrier to the first chemical step by 3.0 kJ/mol. Assuming that hydrolysis of the glycosyl-enzyme is rate-determining, the ratio of kcat values indicates that galactosylation also increases the rate of this step, albeit by a modest 1.4-fold. Notably, PpXG5 exhibited a comparatively low specificity constant for GGGG-CNP, which can be considered a XXXG-CNP homolog with all branching residues removed (Table 3); the kcat/Km value for this substrate was 85-fold lower than for XXXG-CNP and 280-fold lower than for XLLG-CNP. The specificity of PpXG5 is discussed below in light of the three-dimensional structure of the enzyme.

Three-dimensional Structure of PpXG5—The structure of PpXG5 was solved in a native form with data to 1.40 Å resolution. There are two molecules in the asymmetric unit, which are essentially identical. The chain can be traced continuously from residue 33 to 395 in the electron density. It should be noted that the residue numbering corresponds to the protein including a signal peptide, but this was cleaved during the gene expression (as judged by mass spectrometry, which gave an m/z consistent with a mass of ~40,620 Da). PpXG5 displays a (beta/{alpha})8 barrel fold, as is typical of family 5 and clan GH-A enzymes. An open groove runs across the surface of the whole protein, which constitutes the substrate binding subsites (Fig. 5A). Structural similarity searches using the SSM server (62) reveal, not surprisingly, close similarities to other GH-A clan family GH5 enzymes, notably a cellulase from C. cellulolyticum (CelCCA) (47), which has 33% identity and a P-score of 21.8 (corresponding to an r.m.s. deviation of 1.28 Å for 316 aligned C{alpha} positions).

Catalysis by family 5 enzymes occurs with retention of anomeric configuration, which involves a double displacement mechanism and goes via a covalent glycosyl-enzyme intermediate (63). Two carboxylate-containing residues are involved, one that acts as a nucleophile to attack at the anomeric center and another that acts as an acid/base residue to protonate the glycosidic oxygen during the first step of the mechanism and deprotonate a water molecule during the second step. These catalytic residues have been identified as Glu182 (acid/base) and Glu323 (nucleophile) in PpXG5 by analogy with other GH5 enzymes.

Ligand Complex of PpXG5—To determine if crystallization could be used to screen for ligand specificity, PpXG5 was crystallized in the presence of mixed xyloglucan oligosaccharides (2:1:3:3 mixture of XXXG/XLXG/XXLG/XLLG) to obtain structural information on the interactions made with the enzyme. Data on the substrate complex were collected to 1.95 Å resolution. The crystals grew in a different space group to the native crystals, and there is one molecule in the asymmetric unit. The chain can be traced continuously from residue 37 to 395. The native structure and the complex superimpose well, with a r.m.s. deviation of 0.5 Å for the C{alpha} atoms.

There is well defined electron density for a number of sugar rings in the active site of PpXG5, which corresponds to a molecule of XXLG bound in the minus subsites (Fig. 5B). There are four beta-1,4-glucose moieties in subsites -1, -2, -3, and -4, two {alpha}-1,6 xylose residues branched from the glucose moieties in subsites -2 and -3 (the xylose residue that must be present in the -4 subsite is too disordered to be observed in the electron density), and a beta-1,2-linked galactoside linked to the xylose in the -2 subsite. No electron density can be observed for a similarly linked galactose residue in the -3 subsite (which would give XLLG, a molecule also present in the mixture of oligosaccharides co-crystallized with PpXG5).


Figure 6
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  		FIGURE 6.
Schematic diagram of the interactions of PpXG5 with XXLG.

 
The hydroxyl group at C-1 of the glucose residue in the -1 subsite hydrogen-bonds with both the acid/base (Glu182) and nucleophile (Glu323) residues; the C-2 hydroxyl interacts with Glu323, Asn181, and His131, and the C-3 hydroxyl also hydrogenbonds with His131. Asn363 interacts with both of the hydroxyl groups at C-2 and C-3 of the glucose moiety in the -2 subsite, and the C-3 hydroxyl also hydrogen-bonds with Asn50. The xylose residue in the -2 subsite hydrogen-bonds with Ser137. The sugars in the -3 and -4 subsites make no hydrogen bond interactions with the enzyme but only with solvent molecules. There are a number of hydrophobic interactions between aromatic residues and the faces of the sugars, including Trp361 (with glucose in the -1 subsite), Tyr135 (with xylose in the -2 subsite), Trp65 (with glucose in the -3 subsite), and His365 (with xylose in the -3 subsite). Interactions are shown in Fig. 6.

Kinetic Analysis of BlXG12—Of the polysaccharide substrates tested (Table 2), BlXG12 demonstrated highest activity toward xyloglucan, but also showed significant activity toward carboxymethyl cellulose, konjac glucomannan, and barley beta-glucan (Table 3). The pH dependence of the hydrolysis of xyloglucan by BlXG12 was bell-shaped (data not shown), with apparent kinetic pKa values of 4.1 ± 0.1 and 7.8 ± 0.1. Size exclusion chromatography and HPAEC-PAD indicated that the hydrolysis of tamarind xyloglucan by BlXG12 occurred in an endolytic fashion to produce a limit digest composed of a mixture of XXXG, XLXG, XXLG, and XLLG (Fig. 7).


Figure 7
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  		FIGURE 7.
High performance size exclusion (A) and anion exchange (B) chromatographic analysis of the limit digestion of tamarind xyloglucan by BlXG12.

 
BlXG12 was active on the chromogenic substrates GGGG-CNP, XXXG-CNP, XLLG-CNP, and XXXG-pMP (Fig. 8, Table 3). Based upon kcat/Km values, the specificity of this enzyme was inversely related to the degree of substrate branching. BlXG12 hydrolyzed GGGG-CNP to liberate the aglycon with a similar Km value and a ~5-fold greater kcat value compared with XXXG-CNP. XXXG-CNP and XLLG-CNP exhibited similar kcat values, whereas the Km value for the galactosylated substrate was 3.5-fold higher (Table 3). Interestingly, the enzyme was prone to substrate inhibition by XXXG-CNP (Fig. 8A) but not XLLG-CNP (Fig. 8B). Both the substrate inhibition and specificity trends are opposite to those observed for PpXG5. Indeed, the presence of three additional xylosyl units on the Glc4 backbone imposes a 4.5 kJ/mol penalty on the first catalytic step, whereas two additional galactosyl units increases the {Delta}{Delta}G value by a further 3.3 kJ/mol, indicating that branching of the glucan chain retards catalysis by BlXG12. Furthermore, the insensitivity of the Km value of XXXG aryl glycosides to the pKa of the aglycon may indicate that formation of the glycosyl-enzyme intermediate is rate-limiting for these substrates.


Figure 8
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  		FIGURE 8.
Initial rate kinetics of the BlXG12-catalyzed hydrolysis of XXXG-CNP and XLLG-CNP. A, XXXG-CNP data were fitted incorporating substrate inhibition (solid line) to give the kinetic parameters shown in Table 3; fitting of the standard Michaelis-Menten equation (dashed line) results in a clear misfit. B, XLLG-CNP data fit to the Michaelis-Menten equation.

 


Figure 9
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  		FIGURE 9.
A, three-dimensional protein schematic representation of BlXG12 in complex with XXXG/XX. The protein chain is shown color-ramped from the N terminus (blue) to C terminus (red) with the ligand and the catalytic acid/base Glu243 shown in a ball-and-stick representation. B, observed electron density, interpreted as a single molecule of XXXG bound to the -4 to -1 subsites of the enzyme and XX bound to the +1 and +2 subsites. Both images, drawn with MOLSCRIPT (68)/BOBSCRIPT (69), are in divergent (wall-eyed) stereo representation.

 
Three-dimensional Structure of BlXG12—The structure of the BlXG12 nucleophile mutant (E155Q) was solved with data to 1.78 Å resolution. There are two molecules in the asymmetric unit, which are essentially identical. The chain can be traced continuously from residue 31 to 261 in the electron density. The residue numbering corresponds to the protein including a signal peptide, which was cleaved during the gene expression (as judged by mass spectrometry, which gave an m/z value consistent with a mass of ~25,994 Da). BlXG12 displays a beta-jelly roll fold, as shown by other family 12 (and clan GH-C) enzymes. A cleft runs across the surface of the protein, which constitutes the substrate binding subsites; this cleft appears to be deeper than observed with PpXG5 (Fig. 9A). Structural similarity searches using the SSM server (62) confirms similarities to GH12 family enzymes, with the closest apparent match being the Humicola grisea Cel12A (HgGH12) (64) (which has 23% identity and a P-score of 6.9, corresponding to an r.m.s. deviation of 1.52 Å for 209 matched C{alpha} positions).

Family 12 enzymes, like family 5 enzymes, catalyze with retention of anomeric configuration in a two-step mechanism. BlXG12 and related enzymes do, however, possess an acid/base residue that protonates syn to the pyranoside O-5-C-1 bond, in contrast to family 5 enzymes which are anti-protonators (65). The important catalytic residues in BlXG12 are Glu243 (acid/base residue) and Glu155 (nucleophile residue, which has been mutated during the structural studies described here).

Ligand Complex of BlXG12—BlXG12 E155A was crystallized in the presence of the same xyloglucan oligosaccharide mixture as described for PpXG5, and data were collected to 1.40 Å resolution. Once again, the substrate complex crystallized in a different space group to the unliganded (E155Q) structure, and there is one molecule in the asymmetric unit. The chain can be traced continuously from residue 29 to 261. The unliganded and complex structures superimpose with an r.m.s. deviation of 0.6 Å for the C{alpha} atoms.


Figure 10
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  		FIGURE 10.
Schematic diagram of the interactions of BlXG12 with XXXG/XX.

 
The electron density for the substrate complex of BlXG12 clearly shows a number of sugar rings in both the positive and negative subsites, corresponding to the observation of two molecules of XXXG (Fig. 9B). There are four beta-1,4-glucose moieties in subsites -1, -2, -3, and -4 and two {alpha}-1,6-xylose residues branched from the glucose moieties in subsites -2 and -3 (the xylose residue that must be attached to the glucose in the -4 subsite is too disordered to be observed in the electron density). No electron density can be observed for beta-1,2-linked galactose residues on the xylose residues in either the -2 or -3 subsites. Similarly, there are two beta-1,4-glucose moieties in subsites +1 and +2 and two {alpha}-1,6-xylose residues branched from each of them. There is disordered electron density in the +3 subsite corresponding to the third glucose residue, but this cannot be built with confidence, and neither the likely xylose moiety in the +3 subsite nor the glucose residue in the +4 subsite are observed.

The hydroxyl group at C-1 of the glucose residue in the -1 subsite is observed to mutarotate; both the {alpha}- and beta-anomers interact with Met157, and the beta-anomer also interacts with Asp137 (interactions are shown in Fig. 10). The C-2 hydroxyl group hydrogen-bonds with Trp197, the C-6 hydroxyl group hydrogen-bonds with Trp53 and Glu243 (the acid/base residue), and the endocyclic oxygen also interacts with Glu243. The hydroxyl group at C-2 of the glucose moiety in the -2 subsite interacts with Asn51, and the C-3 hydroxyl interacts with His97; the xylose residue in the -2 subsite only makes interactions with solvent molecules. Neither of the glucose moieties in the -3 and -4 subsites make any hydrogen bond interactions with protein residues. The C-4 hydroxyl group of the xylose residue in the -3 subsite, however, hydrogen-bonds to both Ser36 and the main chain nitrogen of Val52. Hydrophobic interactions are made between Trp139 and the glucose moiety in -1, Trp53 and the glucose moiety in -2, and Trp98 and the glucose moiety in -3.

The sugars bound in the plus subsites of BlXG12 make relatively few interactions with the protein. The hydroxyl groups at C-3 and C-4 of the glucose moiety in the +1 subsite both interact with the acid/base residue (Glu243), and the hydroxyl group at C-2 hydrogen bonds with the main chain carbonyl group of Gly166. The xylose moiety in the +2 subsite hydrogen-bonds with the main chain nitrogen atom of Gly166 and stacks with Phe245. Neither the xylose residue in the +1 subsite nor the glucose moiety in the +2 subsite make any interactions with the enzyme.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
What constitutes a xyloglucanase? Formally one might define a xyloglucanase as an enzyme with a catalytic preference for xyloglucan substrates, as opposed to other glucans. In practice, such a distinction is difficult, since long unsubstituted beta-1,4 glucans are insoluble and hence kinetically intractable. In order to analyze xyloglucan specificity, one therefore performs kinetics on polymeric substrates, typically tamarind xyloglucan and a range of unsubstituted and diverse beta-glucans, which includes both artificial soluble beta-1,4-glucans, such as carboxymethyl cellulose, as well as natural beta-glucans, such as barley beta-glucan (mixed beta-1,3 and beta-1,4 bonds) and glucomannans.


Figure 11
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  		FIGURE 11.
Three-dimensional protein schematic representation of (A) PpXG5 in complex with XXLG (green) superposed with a cellulase from C. cellulolyticum (Protein Data Bank entry 1EDG) (yellow) and (B) BlXG12 in complex with XXXG/XX (green) superposed with an endoglucanase from H. grisea in complex with cellopentaose (Protein Data Bank entry 1UU6) (yellow). The figure was drawn with MOLSCRIPT (68) and is in divergent (wall-eyed) stereo representation.

 
PpXG5 and BlXG12 exemplify the spectrum of enzyme activities that one might appropriately term xyloglucanases. BlXG12 is more active on tamarind xyloglucan than the other polysaccharides tested, yet it is only slightly better on this substrate than on the best artificial substrate, low viscosity carboxymethyl cellulose. On the panel of aryl oligosaccharides examined, BlXG12 clearly prefers a naked glucan chain in the glycon (negative) subsites, and catalysis is impaired in the presence of xyloside and further galactoside substituents. Despite this kinetic preference, BlXG12 does not legislate against galactose substituents, since limit digest analysis gives the full spectrum of possible xyloglucan oligosaccharides (based upon a Glc4 backbone) such that XXLG, XLXG, and XLLG must be accommodated in both negative and positive subsites of the enzyme.

In contrast to the accommodation of side chain sugars by BlXG12 and the partial preference of the enzyme for tamarind xyloglucan, PpXG5 is a significantly "better" and more specific xyloglucanase under the conditions used. Indeed, PpXG5 is active only on the substituted polymer and not on any of the other polysaccharides tested, and this activity is extremely high compared with BlXG12. Furthermore, PpXG5 favors xyloglucan oligosaccharides in its negative subsites such that, for good leaving groups, binding and formation of the covalent intermediate are extremely rapid, and deglycosylation is most likely rate-limiting. Furthermore, the galactoside moieties are harnessed productively by the enzyme, as reflected in the 4-fold better catalytic efficiencies on XLLG-CNP versus XXXG-CNP.

This span of different xyloglucan specificities from toleration and partial harnessing by BlXG12 through to absolute specificity and harnessing of extended substituents by PpXG5 is well reflected in their respective three-dimensional structures. For studies of both enzymes in complex with ligand, we intentionally screened a mixture of oligosaccharides in order to sift the most favored xyloglucan-derived oligosaccharide from the 2:1:3:3 mixture of XXXG/XLXG/XXLG/XLLG. In the case of BlXG12, this yielded a -4XXXG-1 +1XX+2 complex, reflecting binding of a minor component of the mixture, but entirely consistent with the kinetic preference for the XXXG-aryl substrate over the galactosylated substrates. Strong positive subsite binding may also be reflected in the substrate inhibition observed for this enzyme (as is the case with, for example, Humicola insolens Cel7B (67)). In the case of PpXG5, the same experiment yielded XXLG bound in the -4 to -1 subsites, consistent with both the importance of these negative subsites to formation of the covalent intermediate and the kinetic preference for galactosyl moieties.

A comparison of PpXG5 and BlXG12 with other members of their respective families gives an insight into what allows them to act on xyloglucan-derived substrates or indeed prevents other enzymes from having this capacity. Inspection of the active site overlap of the closest structural homolog to PpXG5, CelCCA from C. cellulolyticum (47) (Fig. 11A), as well as primary sequence alignments of family 5 members, reveals that Trp361 and Trp65, which stack with glucose residues in the -1 and -3 subsites, respectively, are conserved among GH5 members. However, Ser137, which interacts with the C-3 hydroxyl group of the xylose residue in the -2 subsite, and Tyr135, which stacks with the same xylose residue, are found on a loop that is positioned differently in CelCCA and other GH5 members, such as Cel5A (a cellulase from Bacillus agaradhaerans (66)). Primary sequence alignments of about 20 family GH5 open reading frames shows that PpXG5 and the close homologs from Bacillus sp. BP-23 (55) and Paenibacillus sp. KM21 (54) have either a serine or threonine residue at this position, which is in a conserved region with the motif GDG(F/Y)(H/N)(S/T)(I/V), which is not apparent in any of the other family 5 sequences. Likewise, His365, which stacks with the xylose residue in the -3 subsite, appears to be in a conserved YWDNG(H/F) motif when compared with the Bacillus sp. BP-23 and Paenibacillus sp. KM21 sequences, but which is missing from other GH5 sequences. The superposition of the PpXG5 structure with CelCCA and Cel5A shows that although the protein backbone for each structure is in a similar position in the region of His365, CelCCA and Cel5A have nonaromatic residues in this position that are not in an orientation to interact with the xylose. As well as PpXG5 possessing residues that promote interactions with xyloglucan-derived substrates, other GH5 members possess residues that are likely to prevent binding of them. For example, His123 (Cel-CCA) or Tyr66 and Leu103 (Cel5A) would clash with the xylose residue in the -2 subsite, Phe42 (CelCCA) or Ser69 (Cel5A) would block the galactose residue in the -2 subsite, and Lys267 (Cel5A) would prevent a xylose binding in the -3 subsite.

Rationalization of the BlGH12 xyloglucanase activity compared with other GH12 members is, however, more difficult. BlGH12 makes no interactions with the xylose residue in the -2 subsite, and superposition with its closest structural homologue, HgGH12 from H. grisea (Protein Data Bank entry 1UU6 (64)) (Fig. 11B), demonstrates that a xylose would not clash with any active site residues. However, Tyr9 of HgGH12 would clash with the xylose in the -3 subsite; in this equivalent position in BlGH12, there is a serine that makes productive hydrogen bond interactions with the xylose. In the positive subsites, the overlap with HgGH12 shows that there is nothing to prevent binding of a xylose residue in the +1 subsite, but Tyr132 and Arg97 would block binding in the +2 subsite; in the equivalent positions, BlGH12 possesses a threonine and glycine, respectively. The lack of information about whether other GH12 members possess xyloglucanase activity makes it difficult to draw conclusions about whether the type of residues at these positions can be used to predict the activity of an individual enzyme.

PpXG5 and BlXG12, together with the recently studied CtXG74 xyloglucanase (41), highlight the diversity of microbial xyloglucan-active hydrolases available in nature. Given the massive importance of biofuels and the potential applications of xyloglucan oligosaccharides, the challenge remains to determine how best to harness this spectrum of activities for optimal applied usage.


   FOOTNOTES
 
* This work was supported by the Swedish Foundation for Strategic Research, the Swedish Research Council, and the Biotechnology and Biological Sciences Research Council. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 These authors contributed equally to this work. Back

2 A Fellow (Rådsforskare) of the Swedish Research Council. To whom correspondence may be addressed. E-mail: harry{at}biotech.kth.se.

3 Recipient of a Royal Society-Wolfson Research Merit Award. To whom correspondence may be addressed. E-mail: davies{at}ysbl.york.ac.uk.

4 Y. Kitago, N. Watanabe, S. Karita, K. Sakka, and I. Tanaka, unpublished data. Back

5 The abbreviations used are: HPLC, high pressure liquid chromatography; HPAEC-PAD, high performance anion exchange chromatography system with pulsed amperometric detection; CNP, 2-chloro-4-nitrophenyl; pMP, 4-methoxylphenyl; BisTris, 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol; r.m.s., root mean square. Back

6 F. M. Ibatullin, M. J. Baumann, and H. Brumer, manuscript in preparation. Back


   ACKNOWLEDGMENTS
 
We thank Martin J. Baumann (KTH Biotechnology) for helpful discussions and assistance with HPLC analysis.



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