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Appl Environ Microbiol. Nov 2011; 77(21): 7541–7550.
PMCID: PMC3209137

Functional Analyses of Multiple Lichenin-Degrading Enzymes from the Rumen Bacterium Ruminococcus albus 8[down-pointing small open triangle]

Abstract

Ruminococcus albus 8 is a fibrolytic ruminal bacterium capable of utilization of various plant cell wall polysaccharides. A bioinformatic analysis of a partial genome sequence of R. albus revealed several putative enzymes likely to hydrolyze glucans, including lichenin, a mixed-linkage polysaccharide of glucose linked together in β-1,3 and β-1,4 glycosidic bonds. In the present study, we demonstrate the capacity of four glycoside hydrolases (GHs), derived from R. albus, to hydrolyze lichenin. Two of the genes encoded GH family 5 enzymes (Ra0453 and Ra2830), one gene encoded a GH family 16 enzyme (Ra0505), and the last gene encoded a GH family 3 enzyme (Ra1595). Each gene was expressed in Escherichia coli, and the recombinant protein was purified to near homogeneity. Upon screening on a wide range of substrates, Ra0453, Ra2830, and Ra0505 displayed different hydrolytic properties, as they released unique product profiles. The Ra1595 protein, predicted to function as a β-glucosidase, preferred cleavage of a nonreducing end glucose when linked by a β-1,3 glycosidic bond to the next glucose residue. The major product of Ra0505 hydrolysis of lichenin was predicted to be a glucotriose that was degraded only by Ra0453 to glucose and cellobiose. Most importantly, the four enzymes functioned synergistically to hydrolyze lichenin to glucose, cellobiose, and cellotriose. This lichenin-degrading enzyme mix should be of utility as an additive to feeds administered to monogastric animals, especially those high in fiber.

INTRODUCTION

The rumen microbiome has evolved efficient mechanisms to degrade plant cell wall polysaccharides into metabolizable energy. Ruminococcus albus is a fibrolytic bacterium found in the rumen of many herbivores (3, 23, 32, 45). This bacterium is capable of efficient fermentation of cellulose as well as hemicellulose for growth. Degradation of cellulose, consisting of glucose monomers polymerized by β-1,4 glycosidic bonds, requires the concerted action of 3 classes of enzymes, endoglucanases (EC 3.2.1.4), exoglucanases (EC 3.2.1.91), and β-glucosidases (EC 3.2.1.21), which can be in a noncomplexed or complexed (cellulosomal) state (2, 9, 13, 24, 29). The hemicellulose xylan is a complex heteropolymer consisting of a β-1,4-linked xylose backbone that may be appended with a variety of substituents and requires a wide array of enzymes for full depolymerization (68). Mixed-linkage β-glucans are composed of glucose monomers that are linked together by β-1,3 and β-1,4 glycosidic bonds. These polymers are involved in maintaining the plant cell wall structure and providing stored energy in the plant order Poales, with members such as barley, wheat, and ryegrass (4, 5). Lichenin, another mixed-linkage β-glucan, is found as a storage polysaccharide in lichens, which are forage for reindeer and caribou during the winter season (1, 19, 25).

Bacterial enzymes specific for lichenin degradation are classified within the glycoside hydrolase (GH) family 16 lichenases (EC 3.2.1.73) and selectively hydrolyze β-1,4 glycosidic bonds of 3-O-substituted glucosyl residues (35). Lichenases have been studied through biochemical assays as well as crystal structures and binding studies, and the residues involved in substrate recognition and hydrolysis in some enzymes have been identified (15, 34, 39).

The enzymes involved in cellulose deconstruction, including endo- and exoglucanases from GH families 5 and 9, interact with and hydrolyze lichenin at nonspecific β-1,4 linkages (21). Laminarin, a β-1,3-linked glucose polymer with β-1,6-linked glucose substituents, is hydrolyzed by GH16 laminarinases (EC 3.2.1.39), which also have the ability to degrade internal β-1,3 linkages within lichenin (10).

Several biochemical studies have been conducted on lichenin-degrading enzymes of ruminal origin; however, only two enzymes of the ruminal cellulolytic bacterium Fibrobacter succinogenes have been extensively characterized at the biochemical level (12, 33, 37, 40). A bioinformatic search of a partial genome of Ruminococcus albus 8 revealed many genes encoding putative glycoside hydrolases. Three cloned endoglu-canases, two from R. albus F-40 and one from R. albus SY3, were capable of hydrolyzing lichenin, but the mechanisms of their activities were not characterized in detail (22, 30, 36). A clearer understanding of the R. albus enzymes used to degrade complex polysaccharides into fermentable sugars will provide mechanistic insights into the role of strains of this bacterium in their natural habitat. In this study, three genes encoding putative lichenin-degrading enzymes and one gene encoding a putative β-glucosidase in R. albus 8 were expressed, and the hydrolytic activities on polysaccharides and oligosaccharides were biochemically characterized. Insights into the biochemical function of each enzyme and how the enzymes contribute in synergy to effectively degrade lichenin into glucose, cellobiose, and cellotriose for subsequent utilization by the organism are discussed.

MATERIALS AND METHODS

Materials.

R. albus 8 genomic DNA was obtained from the Department of Animal Sciences, University of Illinois at Urbana—Champaign (20). Escherichia coli JM109 and E. coli BL21-CodonPlus(DE3) RIPL competent cells and PicoMaxx high-fidelity DNA polymerase were purchased from Stratagene (La Jolla, CA). The pET-46 Ek/LIC vector kit was obtained from Novagen (San Diego, CA). A QIAprep Spin Miniprep kit was obtained from Qiagen (Valencia, CA). The 1,3-1,4-β-gluco-oligosaccharides, cello-oligosaccharides, laminaribiose, lichenin, laminarin, glucomannan, and wheat arabinoxylan (WAX) were purchased from Megazyme (Bray, Ireland). The glucose oxidase reaction reagents were obtained from Pointe Scientific Inc. (Canton, MI). All other reagents were of the highest possible purity and were purchased from Sigma-Aldrich (St. Louis, MO).

Gene cloning, expression, and protein purification.

A search of the partial genome sequence of R. albus 8 yielded more than 10 genes predicted to encode enzymes capable of cleaving β-1,4 or β-1,3 glycosidic bonds. All of these genes have been cloned, expressed, and partially characterized (M. Iakiviak et al., unpublished data). Two genes, designated ra0453 and ra2830, were annotated as putative exoglucanases, ra1595 was predicted to encode a β-glucosidase, and ra0505 was predicted to be a lichenase. The genomic DNA of R. albus 8 was extracted using a DNeasy blood and tissue kit (Qiagen). The primer pairs, listed in Table 1, were used to amplify their target genes by using a PicoMaxx PCR kit (Agilent Technologies) and R. albus 8 genomic DNA as the template. The forward primers were designed to delete the putative signal peptides present in two of the genes. Thus, from Ra0505, the PCR amplification deleted the N-terminally located sequence MKKITALTLAFMTVFSLSACG, and from Ra2830, MLKKIISGVTAAASACTIVLSTASGVIVHDAPAASTVSA was deleted. The signal peptides were predicted using the SignalP (version 3.0) online server (11). Each PCR product was cloned into the pET-46b vector (Ek/LIC; Novagen), and the ligation products were transformed into Escherichia coli JM109 using electroporation. After selection on lysogeny broth (LB) plates supplemented with ampicillin at 100 μg/ml, five colonies were picked and cultured in LB medium containing the same antibiotic at the same concentration. Plasmids were purified from each culture using a QIAprep Spin Miniprep kit (Qiagen). To confirm that the plasmids contain the desired genes, DNA sequencing was performed on the DNA inserts (W. M. Keck Center for Comparative and Functional Genomics).

Table 1.
Primers used in this study

For gene expression, each plasmid containing the correct DNA insert was transformed into E. coli BL21-CodonPlus(DE3) RIPL competent cells using the heat shock method, and cells were grown on LB agar plates supplemented with ampicillin (100 μg/ml) and chloramphenicol (50 μg/ml) for 16 h. For each gene, five colonies were picked and inoculated into LB medium (10 ml) with ampicillin (100 μg/ml) and chloramphenicol (50 μg/ml) and grown for 6 to 8 h at 37°C with vigorous shaking. After the precultures reached saturation, each culture was transferred to fresh LB medium (1 liter) containing ampicillin and chloramphenicol at the same concentrations stated above and grown at 37°C with shaking until the optical density at 600 nm (OD600) reached 0.5. Gene expression was induced by adding isopropyl-β-d-thiogalactopyranoside (IPTG) to a final concentration of 0.1 mM, and the cultures were incubated at 16°C for 16 h.

Cells were collected by centrifugation (5,000 × g, 15 min, 4°C) and resuspended in 30 ml of lysis buffer (50 mM Tris, 300 mM NaCl, pH 7.5). The cell suspensions were lysed using an EmulsiFlex C-3 cell homogenizer from Avestin (Ottawa, Ontario, Canada). The cell debris was separated by centrifugation (20,000 × g, 30 min, 4°C), and the clarified lysate was incubated at 4°C for 1 h with Talon metal affinity resin (Clontech). The bound proteins were eluted from the resin using an elution buffer composed of the lysis buffer supplemented with 150 mM imidazole. Subsequently, the fractions containing the Ra0453, Ra0505, and Ra2830 proteins were subjected to gel filtration (HiLoad 16/60 Superdex 200 prep-grade column; GE Healthcare) on an AKTAxpress fast protein liquid chromatograph (FPLC; GE Healthcare). The proteins were in a buffer composed of 50 mM Tris, 150 mM NaCl, pH 7.5, and the same buffer was used to develop the chromatography. The highly purified protein fractions (based on sodium dodecyl sulfate-polyacrylamide gel electrophoresis [SDS-PAGE]) were dialyzed against a storage buffer (50 mM Tris, 150 mM NaCl, pH 7.5) and stored at 4°C. The R. albus 8 Ra1595 protein was subjected to gel filtration chromatography with a buffer containing 50 mM sodium phosphate, pH 7.5. The eluted proteins were applied to an anion-exchange column (5-ml HiTrap Q HP column; GE Healthcare), with the buffer used for the gel filtration chromatography serving as the equilibration buffer. To elute the bound proteins, a gradient was developed with the equilibration buffer containing NaCl at a 1 M concentration. The highly purified protein fractions were dialyzed against the protein storage buffer described above and stored at 4°C until used. SDS-PAGE was carried out as described elsewhere (26), and the proteins were visualized by staining with Coomassie brilliant blue G-250. The concentration of each purified protein was determined using absorbance spectroscopy by the method described by Gill and von Hippel (16). Briefly, the NanoDrop 1000 apparatus from Thermo Fisher Scientific Inc. (Waltham, MA) was used to measure the protein concentration on the basis of the molecular mass and extinction coefficients of Ra0453 (41.2 kDa), Ra0505 (29.1 kDa), Ra1595 (101.4 kDa), and Ra2830 (63.4 kDa), which were estimated to be 84,020 M−1 cm−1, 56,510 M−1 cm−1, 108,600 M−1 cm−1, and 171,200 M−1 cm−1, respectively.

Hydrolysis of pNP-linked sugars.

Recombinant enzymes were screened on a library of para-nitrophenyl (pNP)-linked glycosides, including pNP-α-l-arabinopyranoside, pNP-α-l-arabinofuranoside, pNP-β-d-fucopyranoside, pNP-α-l-fucopyranoside, pNP-α-d-galactopyranoside, pNP-β-d-galactopyranoside, pNP-α-d-glucopyranoside, pNP-β-d-glucopyranoside, pNP-β-d-maltopyranoside, pNP-α-d-maltopyranoside, pNP-α-d-mannopyranoside, pNP-β-d-mannopyrano-side, pNP-α-l-rhamnopyranoside, pNP-β-d-xylopyranoside, and pNP-β-d-cellobioside. All pNP-linked substrates were purchase from Sigma-Aldrich (St. Louis, MO). R. albus 8 Ra0453, Ra0505, Ra1595, and Ra2830, at 0.5 μM in each case, were incubated with substrate (1 mM) for 30 min in a buffer containing 50 mM sodium phosphate, 150 mM sodium chloride, pH 7.0, at 37°C in a thermostated Synergy II multimode microplate reader from BioTek Instruments Inc. (Winooski, VT). The release of pNP was measured at 400 nm every 31 s using the path-length correction feature to convert the absorbance values recorded to correspond to a 1-cm path length. Absorbance values calculated were converted to concentrations using a predetermined extinction coefficient of 10.601 mM−1 cm−1 for pNP at pH 7.0 and a wavelength of 400 nm. To determine the optimal pH, pNP-cellobioside was dissolved to a final concentration of 1 mM in a series of buffers, including citrate buffer (50 mM sodium citrate, 150 mM sodium chloride, pH 5.0, 5.5, 6.0), phosphate buffer (50 mM sodium phosphate, 150 mM sodium chloride, pH 6.0, 6.5, 7.0, 7.5, 8.0), and bicine buffer (50 mM bicine, 150 mM sodium chloride, pH 8.0, 8.5, 9.0). The specific activities of Ra0453 (6 nM) and Ra2830 (100 nM) were calculated at each pH using an extinction coefficient of pNP determined for the corresponding buffer. The enzymes were at a concentration that provided linear release of pNP. By comparing relative activity at each pH, buffer A (50 mM sodium phosphate, 150 mM sodium chloride, pH 6.5) was determined to be the optimal buffer. Therefore, all subsequent reactions were carried out in buffer A. The specific activities of Ra1595 were determined for pNP-β-d-glucopyranoside (1 mM) and pNP-β-d-cellobioside (1 mM), using enzyme concentrations of 40 nM and 100 nM, respectively. The concentration of pNP released was calculated using a predetermined extinction coefficient of 4.0033 mM−1 cm−1 for pNP at pH 6.5.

Hydrolysis of polysaccharides.

Hydrolysis of natural substrates, including glucomannan, lichenin, laminarin, and carboxymethyl cellulose (CMC), present at 0.5% (wt/vol), by three of the proteins (Ra0453, Ra0505, Ra2830) was performed at 37°C at concentrations that resulted in a linear release of products. Avicel cellulose, phosphoric acid swollen cellulose (PASC), and Whatman filter paper (WFP) at concentrations of 2% (wt/vol) were incubated with Ra0453 and Ra2830 (1 μM) at 37°C for 16 h. The phosphoric acid swollen cellulose was prepared as described elsewhere (44). The release of reducing sugars was detected using the para-hydroxy benzoic acid hydrazide (pHBAH) assay as described by Lever (27). The specific activities of the enzymes with each polysaccharide as substrate were determined on the basis of the amount of reducing ends release by each enzyme per unit time. Cellobiose was used to derive a standard curve for the pHBAH assay. To examine the products of hydrolysis, polysaccharides incubated with enzymes were separated using thin-layer chromatography (TLC) with a mobile phase composed of n-butanol, acetic acid, and water (10:5:1) and visualized by spraying with methanolic orcinol and heating at 75°C for 15 min. The products of hydrolysis were also separated using high-performance anion-exchange chromatography (HPAEC) with a System Gold high-performance liquid chromatograph (HPLC) instrument from Beckman Coulter (Fullerton, CA) fitted with a CarboPac PA1 guard column (4 by 50 mm) and a CarboPac PA1 analytical column (4 by 250 mm) from Dionex Corporation (Sunnyvale, CA). The eluted saccharides were then detected by HPAEC with pulse amperometric detection (PAD) with a pulsed model 5040 amperometric analytical cell and a Coulochem III electrochemical detector from ESA Biosciences (Chelmsford, MA) as described elsewhere (7). To identify and quantify the mono- and oligo-saccharides produced from enzymatically degraded polysaccharides, peak retention times and peak areas from the chromatographs of samples were compared to those of commercially available saccharides analyzed as standards. For time course hydrolysis and determination of synergistic interactions of enzymatic activity on lichenin, the substrate under investigation at 0.5% (wt/vol) was digested with individual enzymes or different mixtures of Ra0453, Ra0505, Ra1595, and Ra2830 present in solution at 0.5 μM, 0.05 μM, 0.1 μM, and 0.5 μM, respectively.

Hydrolysis of gluco-oligosaccharides.

Enzymatic hydrolysis of cello- and β-1,3–β-1,4-gluco-oligosaccharides was carried out with a 10 mM substrate concentration at 37°C. To determine the specific activities of Ra1595 on cellobiose and laminaribiose, the enzyme was added at concentrations that provided a linear release of product. Glucose production was measured with a glucose oxidase reagent set purchased from a commercial vendor (Pointe Scientific) and used according to the instructions of the manufacturer. The hydrolyzed sample and standards with various concentrations of glucose (5 μl) were added to the preheated reagent (400 μl), and the mixture was incubated at 37°C for 5 min in the thermostated Synergy II multimode microplate reader. The absorbance was measured at 510 nm, and the products released in the reaction mixture were estimated using a standard curve generated with known glucose concentrations. Hydrolysis products of cello- and β-1,3–β-1,4-gluco-oligosaccharides were separated and visualized with thin-layer chromatography as described above. The time course of cellohexaose hydrolysis by Ra0453 and Ra2830 was determined to gain insight into their mode of hydrolysis of cellulosic substrates. The substrate (10 mM) was incubated with Ra0453 or Ra2830 at concentrations of 2 μM and 75 nM, respectively. The reaction products were then separated and detected using HPEAC-PAD as described above. Since initial screening showed that Ra0505 and Ra1595 lack cellulolytic activity, their hydrolysis of cellulosic substrates was not further investigated.

RESULTS

Domain analysis of four lichenin-degrading enzymes from R. albus 8.

A bioinformatics approach was used to identify glycoside hydrolase (GH) genes predicted to encode polysaccharide-degrading enzymes in a partial genome sequence of Ruminococcus albus 8 (GenBank accession no. NZ_ADKM00000000; K. E. Nelson et al., unpublished data). A myriad of genes coding for putative glycoside hydrolases that target cellulose were identified and expressed. Two genes encoding GH5 enzymes (Ra0453 and R2830) were selected for further biochemical characterization on the basis of their annotation as putative exoglucanases, an activity critical to cellulose degradation. Each gene was expressed with an N-terminal hexahistidine tag (6-His tag) in E. coli BL-21 CodonPlus RIPL cells to facilitate purification of the gene product. The two proteins degraded lichenin during initial screenings of multiple substrates. Therefore, other putative GH genes (those for Ra0505 and Ra1595) encoding proteins likely to target lichenin were identified, cloned, and expressed for protein purification and functional characterization in combination with the genes for Ra0453 and Ra2830. The objective here was to determine whether these enzymes functioned synergistically to degrade lichenin. The GenBank accession numbers of the genes for Ra0453, Ra0505, Ra1595, and Ra2830 are EGC02962, EGC02853, EGC01435, and EGC04285, respectively.

In Fig. 1A, the domain architectures, as determined by use of the Pfam database (http://pfam.sanger.ac.uk/), of the four cloned enzymes are presented, with arrows designating the locations of primers. Two of the proteins (Ra2830 and Ra0505) were predicted to possess signal peptides. Therefore, the forward PCR primers were designed to remove the signal peptides to ensure accumulation of the recombinant gene products in the E. coli cells. Whereas Ra0453 is composed of only a GH5 domain, the GH5 module of Ra2830 was flanked by an N-terminal signal peptide and a C-terminal dockerin-like domain. The two enzymes are classified within the same family; however, their GH5 catalytic domains shared only 14% identity. Ra0505, which also has an N-terminal signal peptide, contained a GH16 catalytic domain. The GH16 module is known to occur in glycoside hydrolases with lichenase (EC 3.2.1.73), laminarinase (EC 3.2.1.6), agarase (EC 3.2.1.81), or xyloglucan:xyloglucosyl transferase (EC 2.4.1.207) activity. The R. albus 8 Ra1595 possessed the two domains known to occur in GH family 3 proteins, which are usually β-glucosidases (EC 3.2.1.21) or β-xylosidases (EC 3.2.1.37). An interesting feature of Ra1595 is that its C-terminal and N-terminal domains are usually located at the N termini and C termini, respectively, of members of this GH family. The linker region between the two domains (GH3-C and GH3-N) also appears to be longer than that of other GH3 proteins. Whereas the linker regions of other GH3 proteins examined were approximately 65 to 100 amino acid residues long (7), Ra1595 has a linker of 225 amino acid residues.

Fig. 1.
Cloning of 4 glycoside hydrolases from Ruminococcus albus 8 and characterization of their substrate specificities. (A) Domain architecture of the cloned glycoside hydrolases from R. albus 8. Arrows designate the locations of the primers within the gene ...

Analyses of purified lichenin-degrading enzymes.

The R. albus Ra0453, Ra2830, Ra0505, and Ra1595 were purified to near homogeneity (Fig. 1B) and stored at 4°C until used. The predicted molecular masses of the four proteins (Ra0453, 41.2 kDa; Ra2830, 63.4 kDa; Ra0505, 29.1 kDa; Ra1595, 101.4 kDa) were similar to the values estimated by SDS-PAGE. The purified enzymes were screened for activity on a wide range of polymeric substrates by incubating each enzyme (0.5 μM) with individual substrates (0.5%, wt/vol) for 16 h (data not shown). Specific activities were determined with a reducing sugar assay for substrates that were degraded by the enzymes (Fig. 1C). Despite their low amino acid sequence identity, Ra0453 and Ra2830 hydrolyzed the same polysaccharides, although with different specific activities. The highest specific activity of Ra0453 was with lichenin as a substrate, followed by glucomannan, WAX, and CMC. As shown in the results for Ra0453, the activity on the soluble mixed-linkage β-glucan (lichenin) was approximately 1,000 times higher than that on CMC. The R. albus Ra2830 protein also had the highest specific activity on lichenin, and the activities on CMC and glucomannan, which were similar, were about 10 times lower than the activity on lichenin. For this enzyme, the activity on WAX was very low, as was observed for Ra0453. Very large amounts of reducing ends were released from lichenin by Ra0505 at a very low enzyme concentration. The specific activity of this enzyme on lichenin was 50- and 100-fold higher than the activities of Ra0453 and Ra2830, respectively. Whereas Ra0505 did not have hydrolytic activity on β-1,4-linked glucans (based on CMC as a substrate), it hydrolyzed laminarin, a β-1,3-linked glucan, at a low rate. There was also release of a small amount of reducing ends from WAX at a rate comparable to the rates for Ra0453 and Ra2830. The GH3 enzyme, Ra1595, was able to degrade pNP-glucoside and pNP-cellobioside, but the hydrolysis of cellobiose was much lower, i.e., approximately 18-fold lower than the hydrolysis of pNP-glucoside. By using a glucose oxidase-coupled assay, it was determined that laminaribiose, composed of two glucose monomers linked by a β-1,3 glycosidic bond, was degraded 70 times faster than cellobiose, indicating that this enzyme prefers to cleave β-1,3 glycosidic bonds.

HPLC analysis of end products from polysaccharide degradation.

To determine whether differences in end products were displayed during hydrolysis of polysaccharide substrates by Ra0453, Ra2830, and Ra0505, the individual enzymes were incubated with the different polysaccharides overnight, and the soluble products were analyzed by HPAEC-PAD (Fig. 2). Comparison of Ra0453 and Ra2830 activity on cellulosic substrates (Fig. 2A and B) demonstrated that Ra0453 produces a small amount of cellobiose, whereas Ra2830 released much larger amounts of cellobiose, as well as produced some cellotriose and glucose. This finding shows that although each of the two enzymes contains a GH5 module, their substrate recognition and catalytic properties are different. The hydrolysis of konjac glucomannan by both GH5 enzymes resulted in glucose and many oligosaccharides with identities that are not easily resolved (Fig. 2C).

Fig. 2.
Comparison of products of hydrolysis from polysaccharides incubated with Ra0453, Ra0505, or Ra2830. Chromatographs of products of hydrolysis from Avicel (A), phosphoric acid swollen cellulose (PASC) (B), glucomannan (C), and lichenin (D). Enzymatic degradation ...

The products of lichenin degradation by Ra0453, Ra2830, and Ra0505 showed interesting differences (Fig. 2D). The R. albus Ra0453 enzyme seems to produce mainly glucose, cellobiose, cellotriose, and a fourth peak unresolved because it represents multiple peaks. This pattern is different from the end products profile of Ra2830, which contained a variety of products, including glucose, cellobiose, laminaribiose, cellotriose, a β-1,3-1,4-glucotriose (Li3A), and an unidentified product, which we labeled unidentified oligosaccharide. The GH16 enzyme, Ra0505, mainly produced the latter unidentified oligosaccharide (G3, as in Ra2830 in Fig. 2D), in addition to smaller proportions of cellobiose and Li3A.

Hydrolysis of cello-oligosaccharides.

The GH5 proteins Ra0453 and Ra2830 are both able to hydrolyze cellulosic substrates, although they differ in the rate and extent of degradation. An experiment using cello-oligosaccharides of different lengths was carried out to determine the capacity to release smaller products. Both enzymes were able to hydrolyze cello-oligosaccharides of degrees of polymerization greater than 3. In addition, Ra0453 degraded cellotriose (Fig. 3Ai, lane 5), whereas no products were observed with Ra2830 incubated with this substrate (Fig. 3Aii, lane 5). On the basis of the results of both HPAEC-PAD and TLC analyses, the major product of oligosaccharide degradation from both enzymes is cellobiose, which has been shown to be a preferred substrate of R. albus 7 (38), a relative of the bacterium under study.

Fig. 3.
Activities of GH5 enzymes with cello-oligosaccharides and cellulosic substrates. (A) Products released from cello-oligosaccharides of degrees of polymerization of from 2 to 6. Substrates were incubated with and without Ra0453 (i) or Ra2830 (ii). Oligosaccharides ...

To further analyze the action by which the enzymes hydrolyze longer cello-oligosaccharides, they were incubated with cellohexaose for a time course analysis. The products were analyzed by HPAEC-PAD (Fig. 3B). The R. albus 8 Ra0453 mostly cleaved cellobiose from cellohexaose to produce cellotetraose, which was then hydrolyzed into two cellobiose units (Fig. 3Bi). This pattern is suggestive of an exocellulolytic mode of action, as cellobiose constitutes the repeating unit in cellulose. Conversely, Ra2830 displayed an endocellulolytic mode of action, producing relatively equal amounts of cellobiose, cellotriose, and cellotetraose from cellohexaose initially, with further degradation to shorter products, mostly cellobiose, with time (Fig. 3Bii).

Hydrolysis of β-1,3–β1,4-gluco-oligosaccharides.

All of the cloned enzymes had high activity on lichenin or soluble oligosaccharides that can be produced from lichenin hydrolysis. To obtain further insight into the types of bonds cleaved and the subsite recognition by these enzymes, Ra0453, Ra2830, Ra0505, and Ra1595 (0.5 μM) were incubated with gluco-oligosaccharides that contain various configurations of the β-1,3 and β-1,4 glycosidic bonds (Fig. 4).

Fig. 4.
Degradation of mixed-linkage β-1,3-1,4-gluco-oligosaccharides by Ra0453, Ra2830, Ra0505, and Ra1595. Various β-1,3-1,4-glucooligosaccharides (10 mM) with degrees of polymerization of from 2 to 5 were incubated in the presence or absence ...

The R. albus Ra0453 hydrolyzed β-1,4-1,3-glucotriose (Li3B) into glucose and cellobiose (Fig. 4Ai, lanes 6 and 7), β-1,3-1,4-1,4-glucotetraose (Li4A) into cellobiose (2 glucose units joined by a β-1,4 glycosidic linkage) and laminaribiose (2 glucose units joined by β-1,3 glycosidic linkage), β-1,4-1,4-1,3-glucotetraose (Li4B) into cellobiose and glucose (Fig. 4Ai, lanes 10 and 11), Li4C into cellobiose (Fig. 4Ai, lanes 12 and 13), and β-1,3-1,4-1,4-1,4-glucopentaose (Li5A) into cellobiose and Li3A (Fig. 4Ai, lanes 14 and 15). Upon analyzing the substrates that were extensively degraded (Li3B, Li4B, and Li4C), a β-1,3 linkage is cleaved when a β-1,4 linkage is present between the −1 and −2 subsites (Fig. 4Ci). The production of cellobiose and glucose from Li4B can be explained by the slow hydrolysis of cellotriose (further explained later), which is produced from the initial cleavage of the β-1,3 linkage present in the tetrasaccharide (Fig. 4B).

The R. albus Ra2830 was also able to degrade Li4B, and unlike Ra0453, the products included cellobiose and laminaribiose (Fig. 4Aii, lanes 10 and 11). Other oligosaccharides degraded by Ra2830 include Li3B and Li4C (Fig. 4Aii), which were cleaved at the β-1,3 glycosidic bond. Hydrolysis of this linkage was incomplete after 16 h, indicating that these are not preferred substrates.

The R. albus Ra0505 was able to degrade Li4A, Li4C, and Li5A by hydrolyzing a β-1,4 linkage when a β-1,3 linkage is present between the −1 and −2 subsites (Fig. 4Ciii). This activity was evident in oligosaccharides with a degree of polymerization greater than 3 (Fig. 4Aiii).

The R. albus Ra1595 degraded laminaribiose rapidly. In contrast, cellobiose was degraded at a much lower rate (Fig. 1C). These findings indicated a preference for hydrolysis of the β-1,3 glycosidic bond. The experiment with the mixed-linkage cello-oligosaccharides as substrates (Fig. 4Aiv) suggested that Ra1595 preferentially releases the glucose molecule in a β-1,3 glycosidic bond at the nonreducing end. Hydrolysis of Li3A produced glucose and cellobiose, and hydrolysis of Li4A produced glucose and cellotriose with a small amount of cellobiose (Fig. 4Aiv). When the nonreducing end β-1,3-linked glucose molecule is cleaved, the cello-oligosaccharide intermediate accumulates in the solution. However, Li3B was partially degraded into glucose with no accumulation of a disaccharide. This hydrolysis pattern can be attributed to the slow release of the β-1,4-linked glucose present at the nonreducing end of the chain and then rapid hydrolysis of the laminaribiose intermediate that is linked by a β-1,3 glycosidic bond.

Hydrolysis of lichenin by Ra0453, Ra2830, and Ra0505.

The R. albus 8 Ra0453, Ra2830, and Ra0505 were shown to hydrolyze several polysaccharide substrates (Fig. 1 and and2),2), with the highest activities displayed on lichenin. To examine the products released by each of the enzymes during hydrolysis of lichenin, a time course for hydrolysis of this substrate was performed. In Fig. 5A, representative chromatograms show a timed release of products from lichenin hydrolysis, as analyzed by HPAEC-PAD. The concentrations of the products that matched the elution of standards were calculated and are plotted in Fig. 5B. The R. albus Ra0453 (50 nM) hydrolyzed lichenin within 2 h into cellotriose, cellobiose, and glucose (Fig. 5A, Ra0453). Upon continued incubation, the cellotriose was converted to glucose and cellobiose. At the same enzyme concentration as Ra0453, Ra2830 degraded lichenin at a lower rate and released a mixture of products simultaneously (Fig. 5A and B, Ra2830), and the end product profiles remained the same over a period of 12 h, indicating that there are no intermediate oligosaccharides that are further degraded with time. The Ra0505 of R. albus 8 was incubated with lichenin at 5 nM, which is 10 times lower than the concentrations of the other two enzymes; however, the degradation of lichenin was complete within 2 h. The major product was the unidentified oligosaccharide, and there were no detectable intermediate sugars.

Fig. 5.
Hydrolysis of lichenin by Ra0453, Ra2830, and Ra0505 with time. Lichenin at a final concentration of 0.5% (wt/vol) was incubated for various lengths of time with Ra0453, Ra2830, or Ra0505 each at a final concentration of 0.5 μM. (A) Identification ...

Synergistic hydrolysis of lichenin by Ra0453, Ra2830, Ra0505, and Ra1595.

The R. albus 8 Ra1595 yielded almost no detectable products from lichenin, on the basis of HPLC analysis (Fig. 6A). However, since it degraded oligosaccharides derived from lichenin, it was analyzed together with the polysaccharide-degrading enzymes to determine if combinations of the four enzymes will synergistically release products. All possible combinations of enzyme mixtures were investigated, and the soluble sugars released were analyzed by HPAEC-PAD. Short-term hydrolysis (Fig. 6), performed for only 1 min, showed pronounced release of cellobiose and cellotriose in mixtures containing Ra0453. In order to quantify the degree of synergy (DOS), the following formula was used: DOS = [total sugars released from enzyme mixture]/Σ([total sugars released from incubation with individual enzymes]). A majority of the mixtures showed some amount of synergistic activity. Combinations of two or three enzymes lacking Ra2830 showed a lower DOS than mixtures containing Ra2830. When Ra2830 is coincubated with Ra1595 on lichenin, there is a 4-fold increase in the concentration of glucose released, leading to a DOS of approximately 1.5. Mixtures containing Ra0453 were able to release larger amounts of glucose and cello-oligosaccharides, and the unidentified oligosaccharide was not detected. The mixture containing all of the enzymes released the most glucose, cellobiose, and cellotriose, which corresponded to approximately 85% conversion of the substrate in the reaction mixture into soluble sugars. After 12 h of hydrolysis (see Fig. S1 in the supplemental material), mixtures containing Ra0453 were able to release only glucose, cellobiose, and cellotriose, while mixtures lacking Ra0453 contained large amounts of the unidentified oligosaccharide (G3 in Fig. S1 in the supplemental material). The unidentified oligosaccharide was purified from thin-layer chromatography plates and subjected to hydrolysis by each of the enzymes investigated in the present study, and only Ra0453 hydrolyzed this oligosaccharide to cellobiose and glucose (see Fig. S1B, lane 3, in the supplemental material). Thus, this hydrolytic activity of Ra0453 leads to the absence of the unidentified oligosaccharide in the end products of enzyme mixtures containing Ra0453.

Fig. 6.
Determination of synergistic activities of Ra0453, Ra2830, Ra0505, and Ra1595 during hydrolysis of lichenin. All possible combinations of the four enzymes Ra0453, Ra2830, Ra0505, and Ra1595 present at final concentrations of 0.5 μM, 0.5 μM, ...

DISCUSSION

In the plant cell wall of several commonly used livestock feeds, β-glucans serve as cross-linking glycans, holding cellulose microfibrils together and potentially shielding them from microbial degradation. However, lichens possess mixed-linkage β-glucans as a storage polysaccharide and are consumed by reindeer and caribou as forage during winter (19, 25). The mixed linkages found in the feed of ruminants have led to evolution or acquisition of diverse glycoside hydrolases to hydrolyze complex polysaccharides, and these enzymes include cellulases from GH families 5 and 9 (21), as well as GH16 lichenin-specific glucanases (35). In a recent metagenomic study, genes for GH family 16 enzymes were found to comprise 1.74% of the total glycoside hydrolase genes in fiber-adherent bacteria of the bovine rumen, providing support for the importance of mixed-linkage glucan hydrolysis (17).

The R. albus 8 Ra0505 effectively hydrolyzes lichenin, and homologs of this enzyme are found in the sequenced genomes of several important ruminal fibrolytic bacteria, including Butyrivibrio proteoclasticus B316, Prevotella ruminicola 23, Ruminococcus flavefaciens FD-1, two homologs in Fibrobacter succinogenes S85, and two homologs in R. albus 7. Interestingly, the genome sequence of P. ruminicola 23 shows a region that may be involved in lichenin utilization. Within this region are genes coding for a TonB-dependent transporter, a GH16 enzyme, a GH5 enzyme, and a GH3 enzyme in an operon-like organization. The glycoside hydrolases in the putative operon exhibited homology to the enzymes characterized in this study, including Ra0505, Ra0453, and Ra1595. The data presented here may, therefore, provide a model for the mechanism by which P. ruminicola 23 degrades and metabolizes mixed-linkage glucans. A lichenase (XynD) from R. flavefaciens 17 has been cloned and biochemically characterized. The polypeptide of the R. flavefaciens 17 XynD is composed of two catalytic domains: a GH11 xylanase and a GH16 lichenase (12). The GH16 domain of XynD and that of Ra0505 share 55% amino acid sequence identity. In contrast to R. flavefaciens strain 17, the sequenced genome of R. flavefaciens strain FD-1 contains a gene coding for a single polypeptide that harbors three tandem repeats of the GH16 module.

The R. albus 8 Ra0453 is able to efficiently degrade the mixed-linkage glucan into glucose, cellobiose, and cellotriose, mostly products that lack β-1,3 linkages. Several characterized GH5 enzymes possess hydrolytic activity on lichenin. These enzymes have high activity on cellulosic substrates and are likely to hydrolyze the β-1,4 linkages within mixed-linkage glucans and not the β-1,3 linkages (14, 33, 41). The results presented from this study show that Ra0453 is able to hydrolyze the β-1,3 linkages present in β-1,3-1,4-glucans and has no activity on β-1,3-glucans such as laminarin (data not shown). It appears that a β-1,4 linkage present between the −1 and −2 subsites is required by Ra0453 for the hydrolysis of the β-1,3 glycosidic bond (Fig. 4Ci). Although Ra0453 is capable of hydrolyzing the mixed-linkage polysaccharide, this enzyme is predicted to be intracellularly located due to the lack of a signal peptide, and therefore, it may degrade the products of lichenin breakdown after transport into the cell. This hypothesis is supported by the release of the unidentified oligosaccharide product of lichenin hydrolysis by Ra2830 and Ra0505 (enzymes with signal peptides), which is converted to glucose and cellobiose by Ra0453.

There is potential for Ra0453 to be used as feed additive for diets made with barley grain. Previous studies have shown that inclusion of a β-1,3-1,4-glucanase improved the nutritive value of barley-based broiler diets (42). In contrast to other licheninases, the hydrolysis of mixed-linkage glucans by Ra0453 releases a fair amount of glucose with prolonged incubation, and inclusion in feeds may provide additional nutritional benefits, especially to monogastric animals.

There have been two previous studies conducted on β-glucosidases from R. albus strains F-40 and AR67 (31, 43). In the study conducted by Ware et al., the GH3 protein had greater hydrolytic activity on β-1,3-linked laminaribiose and laminarin than the β-1,4-linked cellobiose and cello-oligomers (43). The study performed by Ohmiya et al. characterized a β-glucosidase (31) that shares 79% amino acid sequence identity with Ra1595. This enzyme was able to degrade pNP-β-d-glucoside within 2 h; however, hydrolysis of cellobiose was incomplete after 15 h. In our study, Ra1595 exhibited a similar activity since it was able to hydrolyze pNP-β-d-glucoside faster than cellobiose (Fig. 1Civ). We have shown that Ra1595 preferentially degrades laminaribiose, and therefore, it can be classified as a β-1,3-glucosidase. The products from Ra2830-catalyzed hydrolysis of lichenin include laminaribiose and Li3A, which are suitable substrates for Ra1595. As expected, these two enzymes acted synergistically to release more glucose.

On the basis of the results from the present study, a model for utilization of lichenin by R. albus 8 likely involves secretion of Ra2830 and Ra0505 into the extracellular space to degrade this complex plant cell wall polysaccharide. The two enzymes hydrolyze the β-glucans into a variety of oligosaccharides that are transported into the cell as substrates for further degradation by the other enzymes (Ra0453 and Ra1595) predicted to be intracellularly located. The β-glucosidase Ra1595 degrades laminaribiose and Li3A (produced by Ra2830) into glucose and cellobiose. The R. albus 8 Ra0453 degrades the unidentified oligosaccharide (produced by Ra0505 and Ra2830) into glucose and cellobiose, and these products correspond to the final products seen in the mixture of all four enzymes (see Fig. S1 in the supplemental material). The genome of R. albus 8 codes for a putative cellobiose phosphorylase that likely catalyzes the phosphorolysis of cellobiose into glucose and glucose-1-phosphate, which can then enter glycolysis (18, 28). Future studies on enzyme localization, especially for Ra0453, and gene expression profiles will be required to validate this model.

Supplementary Material

Supplemental Material:

ACKNOWLEDGMENTS

This research was supported by a grant from the Energy Biosciences Institute. Michael Iakiviak was supported by a Nesheim Fellowship in Nutrition from the Department of Animal Sciences, University of Illinois.

We thank Shosuke Yoshida, Yejun Han, Dylan Dodd, Young-Hwan Moon, and Xiaoyun Su for valuable scientific discussions and Anton Evans and Beth Mayer for technical assistance.

Footnotes

Supplemental material for this article may be found at http://aem.asm.org/.

[down-pointing small open triangle]Published ahead of print on 2 September 2011.

REFERENCES

1. Ahmadjian V. 1965. Lichens. Annu. Rev. Microbiol. 19:1–20 [PubMed]
2. Bayer E. A., Belaich J. P., Shoham Y., Lamed R. 2004. The cellulosomes: multienzyme machines for degradation of plant cell wall polysaccharides. Annu. Rev. Microbiol. 58:521–554 [PubMed]
3. Bryant M. P. 1959. Bacterial species of the rumen. Bacteriol. Rev. 23:125–153 [PMC free article] [PubMed]
4. Burton R. A., Fincher G. B. 2009. (1,3;1,4)-β-d-glucans in cell walls of the Poaceae, lower plants, and fungi: a tale of two linkages. Mol. Plant 2:873–882 [PubMed]
5. Chesson A., Gordon A. H., Lomax J. A. 1985. Methylation analysis of mesophyll, epidermis, and fibre cell-walls isolated from the leaves of perennial and Italian ryegrass. Carbohydr. Res. 141:137–147
6. Dodd D., Cann I. K. O. 2009. Enzymatic deconstruction of xylan for biofuel production. Glob. Change Biol. Bioenergy 1:2–17 [PMC free article] [PubMed]
7. Dodd D., Kiyonari S., Mackie R. I., Cann I. K. O. 2010. Functional diversity of four glycoside hydrolase family 3 enzymes from the rumen bacterium Prevotella bryantii B14. J. Bacteriol. 192:2335–2345 [PMC free article] [PubMed]
8. Dodd D., Mackie R. I., Cann I. K. O. 2011. Xylan degradation, a metabolic property shared by rumen and human colonic Bacteroidetes. Mol. Microbiol. 79:292–304 [PubMed]
9. Doi R. H. 2008. Cellulases of mesophilic microorganisms: cellulosome and noncellulosome producers. Ann. N. Y. Acad. Sci. 1125:267–279 [PubMed]
10. Driskill L. E., Bauer M. W., Kelly R. M. 1999. Synergistic interactions among β-laminarinase, β-1,4-glucanase, and β-glucosidase from the hyperthermophilic archaeon Pyrococcus furiosus during hydrolysis of β-1,4-, β-1,3-, and mixed-linked polysaccharides. Biotechnol. Bioeng. 66:51–60 [PubMed]
11. Emanuelsson O., Brunak S., von Heijne G., Nielsen H. 2007. Locating proteins in the cell using TargetP, SignalP and related tools. Nat. Protoc. 2:953–971 [PubMed]
12. Flint H. J., Martin J., McPherson C. A., Daniel A. S., Zhang J.-X. 1993. A bifunctional enzyme, with separate xylanase and β(1,3-1,4)-glucanase domains, encoded by the xynD gene of Ruminococcus flavefaciens. J. Bacteriol. 175:2943–2951 [PMC free article] [PubMed]
13. Fontes C. M. G. A., Gilbert H. J. 2010. Cellulosomes: highly efficient nanomachines designed to deconstruct plant cell wall complex carbohydrates. Annu. Rev. Biochem. 79:655–681 [PubMed]
14. Foong F., Hamamoto T., Shoseyov O., Doi R. H. 1991. Nucleotide sequence and characteristics of endoglucanase gene engB from Clostridium cellulovorans. J. Gen. Microbiol. 137:1729–1736 [PubMed]
15. Gaiser O. J., et al. 2006. Structural basis for the substrate specificity of a Bacillus 1,3-1,4-β-glucanase. J. Mol. Biol. 357:1211–1225 [PubMed]
16. Gill S. C., von Hippel P. H. 1989. Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182:319–326 [PubMed]
17. Hess M., et al. 2011. Metagenomic discovery of biomass-degrading genes and genomes from cow rumen. Science 331:463–467 [PubMed]
18. Hidaka M., et al. 2006. Structural dissection of the reaction mechanism of cellobiose phosphorylase. Biochem. J. 398:37–43 [PMC free article] [PubMed]
19. Holleman D. F., Luick J. R., White R. G. 1979. Lichen intake estimated for reindeer and caribou during winter. J. Wildl. Manage. 43:192–201
20. Hungate R. E., Stack R. J. 1982. Phenylpropanoic acid: growth factor for Ruminococcus albus. Appl. Environ. Microbiol. 44:79–83 [PMC free article] [PubMed]
21. Kanda T., Yatomi H., Makishima S., Amano Y., Nisizawa K. 1989. Substrate specificities of exo- and endo-type cellulases in the hydrolysis of β-(1,3)- and β-(1,4)-mixed d-glucans. J. Biochem. 105:127–132 [PubMed]
22. Kawai S., et al. 1987. Molecular cloning of Ruminococcus albus cellulase gene. Agric. Biol. Chem. 51:59–63
23. Kong Y., He M., McAlister T., Seviour R., Forster R. 2010. Quantitative fluorescence in situ hybridization of microbial communities in the rumens of cattle fed different diets. Appl. Environ. Microbiol. 76:6933–6938 [PMC free article] [PubMed]
24. Kumar R., Singh S., Singh O. V. 2008. Bioconversion of lignocellulosic biomass: biochemical and molecular perspectives. J. Ind. Microbiol. Biotechnol. 35:377–391 [PubMed]
25. Kumpula J. 2001. Winter grazing of reindeer in woodland lichen pasture: effect of lichen availability on the condition of reindeer. Small Rumin. Res. 39:121–130 [PubMed]
26. Laemmli U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685 [PubMed]
27. Lever M. 1972. A new reaction for colorimetric determination of carbohydrates. Anal. Biochem. 47:273–279 [PubMed]
28. Lou J., Dawson K. A., Strobel H. J. 1997. Cellobiose and cellodextrin metabolism by ruminal bacterium Ruminococcus albus. Curr. Microbiol. 35:221–227 [PubMed]
29. Lynd L. R., Weimer P. J., van Zyl W. H., Pretorius I. S. 2002. Microbial cellulose utilization: fundamentals and biotechnology. Microbiol. Mol. Biol. Rev. 66:506–577 [PMC free article] [PubMed]
30. Ohara H., et al. 2000. Sequence of egV and properties of EgV, a Ruminococcus albus endoglucanase containing a dockerin domain. Biosci. Biotechnol. Biochem. 64:80–88 [PubMed]
31. Ohmiya K., Shirai M., Kurachi Y., Shimizu S. 1985. Isolation and properties of β-glucosidase from Ruminococcus albus. J. Bacteriol. 161:432–434 [PMC free article] [PubMed]
32. Orpin C. G., Mathiesen S. D., Greenwood Y., Blix A. S. 1985. Seasonal changes in the ruminal microflora of the high-arctic svalbard reindeer (Rangifer tarandus platyrhynchus). Appl. Environ. Microbiol. 50:144–151 [PMC free article] [PubMed]
33. Palackal N., et al. 2007. A multifunctional hybrid glycosyl hydrolase discovered in a unculturable microbial consortium from ruminant gut. Appl. Microbiol. Biotechnol. 74:113–124 [PubMed]
34. Piotukh K., Serra V., Borriss R., Planas A. 1999. Protein-carbohydrate interactions defining substrate specificity in Bacillus 1,3-1,4-β-d-glucan 4-glucanohydrolase as dissected by mutational analysis. Biochemistry 38:16092–16104 [PubMed]
35. Planas A. 2000. Bacterial 1,3-1,4-β-glucanases: structure, function and protein engineering. Biochim. Biophys. Acta 1543:361–382 [PubMed]
36. Romaniec M. P. M., Davidson K., White B. A., Hazlewood G. P. 1989. Cloning of a Ruminococcus albus endo-β-1,4-glucanase and xylanase genes. Lett. Appl. Microbiol. 9:101–104
37. Taylor E. J., et al. 2005. How family 26 glycoside hydrolases orchestrate catalysis on different polysaccharides: structure and activity of a Clostridium thermocellum lichenase, CtLic26A. J. Biol. Chem. 280:32761–32767 [PubMed]
38. Thurston B., Dawson K. A., Strobel H. J. 1993. Cellobiose versus glucose utilization by ruminal bacterium Ruminococcus albus. Appl. Environ. Microbiol. 59:2631–2637 [PMC free article] [PubMed]
39. Tsai L. C., et al. 2008. Mutational and structural studies of the active-site residues in truncated Fibrobacter succinogenes 1,3-1,4-β-d-glucanase. Acta Crystallogr. D Biol. Crystallogr. 64:1259–1266 [PubMed]
40. Tsai L. C., Shyur L. F., Lee S. H., Lin S. S., Yuan H. S. 2003. Crystal structure of a natural circularly permuted jellyroll protein: 1,3-1,4-β-d-glucanase from Fibrobacter succinogenes. J. Mol. Biol. 330:607–620 [PubMed]
41. Voget S., Steele H. L., Streit W. R. 2006. Characterization of a metagenome-derived halotolerant cellulase. J. Biotechnol. 126:26–36 [PubMed]
42. von Wettstein D., Warner J., Kannangara C. G. 2003. Supplements of transgenic malt or grain containing (1,3-1,4)-β-glucanase increase the nutritive value of barley-based broiler diets to that of maize. Br. Poult. Sci. 44:438–449 [PubMed]
43. Ware C. E., Lachke A. H., Gregg K. 1990. Mode of action and substrate specificity of a purified exo-1,4-β-d-glucosidase cloned from the cellulolytic bacterium Ruminococcus albus AR67. Biochem. Biophys. Res. Commun. 171:777–786 [PubMed]
44. Wood T. M. 1988. Preparation of crystalline, amorphous, and dyed cellulase substrates. Methods Enzymol. 160:19–25
45. Yang S., et al. 2010. Bacterial diversity in the rumen of gayals (Bos frontalis), swamp buffaloes (Bubalus bubalis), and Holstein cow as revealed by cloned 16S rRNA gene sequences. Mol. Biol. Rep. 37:2063–2073 [PubMed]

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