Department of Microbiology, University of Guelph, Guelph, Ontario N1G 2W1, Canada
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Abstract |
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Keywords: catalytic domains/dockerins/family 11 glycosidases/xylanase
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Introduction |
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The xylanases produced by ruminant microorganisms, such as Ruminococcus flavefaciens 17 (Zhang and Flint, 1992) and Neocallismatix patriciarum (Gilbert et al., 1992
; Xue et al., 1992
) are comprised of three domains, two of which are catalytically active. The catalytic domains I and II of N.patriciarum xylanase are essentially homologous and both also belong to family 11 glycosidases. In contrast, the catalytic domains of R.flavefaciens XynA have a very low identity of 19% and, in fact, domain A is assigned to family 10 while domain C is homologous to family 11 enzymes (Zhang and Flint, 1992
). Xylanase C (XynC) produced by the anaerobic, ruminant bacterium Fibrobacter succinogenes S85 (the type strain), has a molecular mass of 63 850 Da and is also comprised of three distinct domains, labelled A, B and C, which are separated by serine-rich linker peptides (Figure 1
). Deletion analysis of the XynC gene indicated that domains A and B function catalytically as xylanases (Paradis et al., 1993
). These two catalytic domains share 43% identity (71% similarity; Figure 2
) and both are members of family 11 glycosidases.
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Materials and methods |
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Ampicillin, L-arabinose, birch wood xylan, D-galactose, D-glucose, 4-morpholinoethanesulfonic acid (MES), 3-(N-morpholino)propanesulfonic acid (MOPS), oat spelt xylan, Triton X-100, Tween 20 and D-xylose were products of Sigma Chemical (St. Louis, MO). Boehringer Mannheim Canada (Laval, Québec) supplied glycine, isopropyl-ß-D-thiogalactoside (IPTG), Pefabloc SC and Tris-base, while xylo-oligomers were obtained from Megazyme (Bray, County Wicklow, Ireland). Reagents for the glutathione-S-transferase purification module (expression vectors, glutathioneSepharose 4B and thrombin protease) were supplied by Pharmacia Biotech (Baie d Urfé, Québec). The bacterial culture medium components tryptone, yeast extract and bacteriological agar were obtaineed from Difco Laboratories (Detroit, MI). All other chemicals were supplied by Fisher Scientific (Nepean, Ont) and were of reagent- or HPLC-grade.
The soluble fraction of oat spelt xylan was prepared as described previously (Bray and Clarke, 1994) from dehydrated oat spelt xylan containing ~10% arabinose and 15% glucose residues as determined by high-pH anion-exchange chromatography (HPAEC) (Clarke et al., 1994).
Cloning and expression of XynC-A and XynC-B
E.coli strain BL-21 [F, ompT, hsdS(rB, mB), gal] containing the construct XynC-B and cloned into the pGEX vector pGEX-4T-1 (Pharmacia Biotech) was kindly provided by C.Forsberg (University of Guelph, Guelph, Ont). The original clone, XynC-BC, containing both domains B and C, was expressed in the recombinant pET-11a in E.coli BL21 (DE3) (Zhu et al., 1994). Domain B was isolated from domain C by PCR amplification using primers coding for the N- and C-terminal ends of domain B which included residues 286529 of XynC (NCIB accession number P35811). The purified DNA product was cloned into the EcoRI and XhoI restriction enzyme site in the multiple cloning region of the pGEX-4T-1 vector and expressed in E.coli BL21.
The gene encoding the A domain of XynC (residues 26288) was amplified from pX14 using primers that incorporated an EcoRI site upstream and a XhoI site downstream from the gene. This construct was subcloned into pGEX-4T-3 (Pharmacia Biotech) to provide pGEX-4T-3XynA. Cells of E.coli DH5 were transformed with pGEX-4T-3XynA for purification of the plasmid and subsequently into E.coli BL21 for expression of the glutathione-S-transferaseXynC-A fusion protein. For the expression of both XynC-A and XynC-B fusion proteins, clones were screened using Congo Red plates incorporating oat spelt xylan and confirmation for the correct inserts was obtained by DNA sequencing analysis.
Bacterial cells from a frozen glycerol stock (70°C) were plated on 2xYTA agar (tryptone 16 g/l, yeast extract 10 g/l, NaCl 5 g/l, agar 15 g/l, pH adjusted to 7.0 with NaOH, supplemented with 100 g/ml ampicillin). Upon growth overnight at 37°C, a single colony was selected for inoculation into an appropriate volume of YTAG [YTA broth supplemented with 2% (w/v) sucrose], which was grown overnight and subsequently inoculated into a larger volume of YTAG for purification. Cells were grown at 30°C with shaking until an absorbance at 600 nm of between 0.5 and 2.0 was reached (~7 h) and then induced for expression of the fusion proteins with the addition of 100 µM of isopropyl-ß-D-thiogalactoside (IPTG) for 3 h. Cells were harvested by centrifugation (16 000 g) for 15 min at 4°C. Supernatants were discarded and cells were frozen at 20°C or processed immediately.
Purification of XynC-A and XynC-B
The same protocol was adopted for the isolation and purfication of both XynC-A and XynC-B enzymes. Cells in 50 µl of ice-cold phosphate-buffered saline (PBS) (140 mM NaCl, 2.7 mM KCl, 10 mM Na2PO4, 1.8 mM KH2PO4, pH 7.3) per ml of culture were sonicated at 150 W for a total of 5 min at 0°C. Triton X-100 was added to the sonicate to a final concentration of 1% and gently mixed at room temperature for 30 min to assist in solubilization of the fusion proteins. The sonicates were isolated by centrifugation (27 000 g) for 15 min at 4°C and to the supernatants was added protease inhibitors (Pefabloc SC) at a final concentration of 0.1 mg/ml.
GlutathioneSepharose 4B, previously equilibrated in PBS, was added to the supernatants at a concentration of 2 ml of resin per 100 ml of sonicate at room temperature with gentle agitation to allow the glutathione-S-transferase component of the fusion proteins to bind to the resin. After 30 min, the resin was separated from the supernatant by centrifugation (500 g) for 5 min at 4°C. The supernatant was discarded and the resin was washed twice with 10 bed volumes of PBS. XynC-A or XynC-B were recovered from the resin by cleavage of the bound fusion protein with the addition of 50 units of thrombin per ml bed volume and incubation at room temperature for 16 h (overnight) with gentle agitation. The suspension was subjected to centrifugation (500 g) at room temperature for 5 min to pellet the Sepharose 4B beads. The eluate was removed, Pefabloc SC at 0.4 mM was added to inhibit further digestion by thrombin and the sample was filtered through a 0.22 µm syringe filter to remove residual Sepharose.
XynC-A and XynC-B were purified to homogeneity by anion-exchange chromatography on Mono Q HR 5/5. The enzymes were applied to the column, previously equilibrated in 10 mM ammonium acetate buffer, pH 7.0, at 1.0 ml/min. Elution was effected with the application of a linear gradient of 10 to 300 mM ammonium acetate buffer over 60 min. Fractions monitored by absorbance at 280 nm were analyzed for activity and protein content and by SDSPAGE. Fractions exhibiting high specific activities and containing apparently homogeneous XynC-A and XynC-B were pooled, lyophilized and resuspended in 50 mM sodium acetate buffer, pH 5.5.
Enzymatic activity and kinetics
Xylanase activity was routinely assayed by monitoring the release of reducing sugars using the arsenomolybdate method of Nelson and Somogyi (Nelson, 1944; Somogyi, 1952). Samples of enzyme (3.264 nM, final concentration) were incubated with the appropriate concentration of substrate in 50 mM sodium acetate buffer, pH 6.5, for between 10 and 20 min at 37°C. Enzyme activity was measured in triplicate using at least two dilutions under all circumstances and was expressed in µmoles of xylose equivalents for the purpose of kinetic analysis, in international units (1 IU = amount of enzyme releasing 1.0 mol of xylose per minute) for specific activity and as percentage maximum activity when appropriate.
The MichaelisMenten parameters Km, Vmax and kcat were determined by incubation of enzyme (3.2 nM) with xylan preparations (at least in triplicate) at final concentrations of 0.115 mg/ml, in 50 mM sodium acetate buffer, pH 6.5, at 37°C and sampled at 0, 1, 2.5, 5, 7.5, 10, 15 and 20 min. Velocities were calculated from linear regions of progress curves and plotted against xylan concentrations. Linear transformation was achieved using the HanesWoolf relationship to depict graphically and evaluate Vmax and Km estimates. Curve-fitting to the MichaelisMenten equation, using non-linear regression analysis and the estimated kinetic constant values, was then conducted to give values for Km and Vmax. Analysis and quantification of reaction products were performed by HPAEC (Bray and Clarke, 1994).
Temperature and pH optima and stability of XynC-B
Unless specified otherwise, temperature and pH optima and stability were assayed in triplicate using 1.01.5% soluble oat spelt xylan as substrate and the NelsonSomogyi reducing sugar assay. Assay results were expressed as percentage maximum activity or as a percentage of an appropriate control. Monotropic buffers (formate, acetate, MES, MOPS, TrisHCl and ethanolamine) at 50 mM concentration and 45°C were used for the pH studies, with the ionic strength being maintained at 50 mM by the addition of an appropriate amount of 1 M KCl. For temperature optima experiments, enzyme (5.4 nM) was incubated with substrate in 50 mM sodium acetate buffer, pH 5.5. Reactions were performed at 20, 30, 35, 40, 45, 50 and 60°C and sampled over time. Temperature stability was evaluated by incubating enzyme at different temperatures (3085°C) for 90 min and then incubating samples with substrate at 45°C to evaluate the remaining activity.
Analytical techniques
Protein concentrations were measured using the BCA protein assay kit from Pierce (Rockford, IL). The Microtiter Plate protocol was used as described by the manufacturer and BSA was used as the standard. Electrospray mass spectrometry was conducted by G.Lajoie (University of Waterloo) using a Micromass Quattro II triple-stage quadrupole mass spectrometer (Micromass, UK) equipped with an electrospray ionization source. SDSPAGE was performed according to the method of Laemmli (1970) using a 12.0% (w/v) polyacrylamide. Gels were stained with either 0.1% (w/v) Coomassie Brilliant Blue R-250 or silver (Bollag and Edelstein, 1991). Zymography (activity gels) was performed as described by Hill (1996) using 12% (w/v) polyacrylamide gels containing 0.1% soluble oat spelt xylan. Western immunoblot analysis was conducted using rabbit antisera to the full XynC enzyme provided by H.Zhu (University of Guelph) and the ammonium sulfate-precipitated polyclonal antibody was used.
Secondary structure predictions were performed using hydrophobic cluster analysis (HCA version 2, Doraine Progiciels Scientifiques, Le Chesnay, France) (Gaboriaud et al., 1987; Lemesle-Varloot et al., 1990
) and the multivariate linear regression combination program (Guermeur et al., 1999
), which combines and analyzes the results of three different prediction programs.
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Results |
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Purification of the pGEX-expressed proteins was conducted using the Pharmacia Biotech GST purification module followed by anion-exchange chromatography on Mono Q. The fusion proteins retained by the glutathioneSepharose affinity column appeared as relatively pure fractions with an apparent molecular weight of 59 kDa (Figure 4A, lane 5), consistent with that expected for the glutathione-S-transferase:XynC-A and -B fusions (28.2 and 27.8 kDa, respectively). The partially purified XynC domains were recovered from the affinity column after thrombin cleavage as protein bands of apparent molecular weight 31 kDa (Figures 3 and 4A
). Although not evident by staining with Coomassie Brilliant Blue, these fractions were contaminated with a number of proteins of both higher and lower molecular weight as detected with silver staining (Figure 4B
, lane 7; note, XynC proteins do not stain with silver). These contaminating proteins were removed by anion-exchange chromatography on Mono Q. A linear gradient of 10300 mM ammonium acetate, pH 7.0, was applied to the column and the XynC derivatives eluted as single peaks at about 35 mM ammonium acetate buffer (data not shown).
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An oat spelt xylan-containing zymogram demonstrated the in situ hydrolysis of substrate by components present in the purification steps (Figure 5). The apparently pure 31 kDa XynC-B successfully hydrolyzed this xylan (lane 6), producing a zone of hydrolysis, as did the 59 kDa fusion protein (lanes 2 and 4). This suggests that fusion of the XynC-B to the glutathione-S-transferase does not impair xylanase activity. Similar results were obtained with the fusion and free forms of XynC-A (data not shown).
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Catalytic parameters of XynC-A and XynC-B
Representative data for the substrate specificity of the two XynC catalytic domains is presented in Table III. As with XynC, XynC-A+ and XynC-BC, both XynC-A and XynC-B catalyzed maximum activity with the xylans from birch wood and oat spelt. The specific activity of both XynC-A and XynC-B against barley ß-glucan was low relative to the xylans, but significantly higher when compared with the other forms of the enzymes acting on this substrate, including XynC-BC. However, like XynC and XynC-A+, XynC-A was essentially inactive against carboxymethylcellulose, whereas XynC-B was only weakly active against this substrate, hydrolyzing it with the same efficiency as XynC-BC.
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Finally, birch wood xylan appeared to serve as a better substrate for both XynC-A and XynC-B than oat spelt xylan, as reflected by the better affinity constants of the enzymes for the former, which typically contains more O-methylglucuronic acid residues and less arabinose side-chains than oat spelt xylan (Clarke, 1997).
Hydrolytic profiles of XynC-A and XynC-B
The major reaction products released by exhaustive hydrolysis of oat spelt xylan by XynC-A and XynC-B were analyzed by HPAEC. Not surprisingly, the enzymes digested this xylan in a manner typical of endo-ß-1,4-xylanases by producing xylooligosaccharides, primarily xylobiose and xylotriose and some xylose (data not shown). These experiments were refined by using purified xylooligosaccharides as substrates and quantifying the reaction products during the course of the reaction. At pH 6.5 and 25°C, 64 nM XynC-B catalyzed the complete hydrolysis of 1 mM xylopentaose (Figure 6) with the production of approximately equal amounts of xylobiose and xylotriose (44.8 and 45.3%, respectively) with the residual as xylose (Table IV
). Likewise, xylotetraose and xylotriose were completely hydrolyzed, but with the production of mostly xylobiose from xylotetraose and both xylobiose and xylose from xylotriose. Xylobiose did not serve as a substrate for XynC-B (Table IV
). These representative data are similar to those obtained previously with XynC-BC (Zhu et al., 1994
) where xylobiose was not hydrolyzed and a significant proportion of xylose was released from the three longer xylooligomers (Table I
). However, as observed with XynC-A+ (Zhu et al., 1994
), the activity of XynC-A is markedly different to that of XynC-B and XynC-BC in that considerably less xylose was released from either xylotetraose or xylopentaose. As with the other forms of the XynC enzymes, XynC-A did not hydrolyze xylobiose (Table IV
).
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The effects of pH and temperature on XynC-B were determined to investigate the influence, if any, of XynC-C on the activity and stability of the catalytic B domain. Preliminary studies indicated that XynC-B was maximally active between pH 4 and 6.5. Formate buffers appeared to enhance activity, as the activity at pH 5.0 was higher in formate than in acetate buffer (data not shown). The activities for the acetate buffer were better than those in MES, MOPS and TrisHCl. In view of its better buffering capacity at pH 5.0, acetate was the buffering system used for further studies. An apparent biphasic response to pH was seen, as shown in Figure 7A (solid curve). The activity was highest between pH 4.5 and 5.5, being optimal at pH 5.0, followed by a slight decrease which remained stable to pH 6.5 and finally a more dramatic loss of activity by pH 7.0. At pH 4.0 and below, there was essentially no detectable activity. The dashed curve in Figure 7A
is representative of the effect of pH on the stability of XynC-B. Essentially, the enzyme activity was stable between pH 5.5 and 7.0, but both below pH 5.0 and above pH 7.5 there was evidence of irreversible inactivation.
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The activity of XynC-B rose sharply between 40 and 50°C and reached a maximum level at ~52°C. Above 55°C, the activity declined rapidly such that at 60°C the enzyme was only 30% as active as at 52°C (Figure 7B, solid line). Activity was also evident at 1840°C, although at a much reduced rate. The enzyme was stable for 90 min when incubated from at least 30 to 65°C. After 90 min at 70°C the enzyme activity was substantially reduced and at 75°C it was insignificant with <10% of the maximum activity reached (Figure 7B
, dashed line).
Secondary structure predictions for XynC domain C
To date, experimentation has failed to identify a function for domain C of XynC. However, the cellulolytic and xylanolytic enzymes of F.succinogenes are thought to form a multi-enzyme complex on the surface of the cells (McGavin and Forsberg, 1988; Gong and Forsberg, 1993
) which may occur in a manner analogous to the well-characterized cellulosomes produced by species of Clostridium (reviewed in Bayer et al., 1998) and R.flavefaciens (Kirby et al., 1997
). Homologous domains to XynC-C do not appear to be present in any other enzyme based on amino acid sequence alignments using various computer algorithms (e.g. BLAST), but hydrophobic cluster analysis revealed a similar pattern of secondary structure when comparing XynC-C with the type I dockerin domains of xylanases and cellulases (Figure 8B
). These apparent structural similarities seem to exist without significant amino acid sequence homologies. For example, there is only 21 and 16% sequence identity (47 and 42% similarity) between XynC-C and the dockerins of R.flavefaciens xylanase B (accession number S51592) and endoglucanase A (accession number Z83304), respectively (Figure 8A
). Moreover, XynC-C lacks the elements of a hypothetical calcium-binding, `F-hand' motif which appears to be present in the R.flavefaciens dockerins. As described by Pagès et al. (1997), this region is characterized by aspartic acid and asparagine residues in positions 1, 3, 5, 9 and 11 with a two-residue species specificity determinant at positions 10 and 11, followed by an
-helix. In the case of the R.flavefaciens EndA and XynB dockerins, the specificity determinant would be Leu and Ala at positions 698/719 and 699/720, respectively (Figure 8A
). However, secondary structure predictions using the multivariate linear regression combination program (Guermeur et al., 1999
) do indicate the presence of an
-helix in the position corresponding to that of the R.flavefaciens dockerins.
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Discussion |
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Of the two isolated catalytic domains, XynC-A appears to be more efficient with one notable exception. Only XynC-B demonstrated activity against carboxymethylcellulose and, as with the other substrates, the separation of domain C from XynC-B significantly enhanced its activity against the soluble cellulose derivative. This difference could be attributed to the previous finding based on a kinetic analysis that domain B has a shorter binding site cleft than domain A (Zhu et al., 1994). It is conceivable that a shorter binding site would enhance the opportunity for the enzyme to associate with the cellulose in regions of less carboxymethyl substitution, which presumably is further enhanced by the absence of domain C.
Another obvious change resulting from the removal of domain C from XynC-BC was the shift in product distribution following activity on xylooligosaccharides, especially xylopentaose. Considerably less xylose was released as a product of the hydrolyses catalyzed by XynC-B than XynC-BC, decreasing to 10% from 22%, respectively, with xylopentaose as substrate. These data suggest that removal of the domain C lessens the multimolecular substrate reactions of XynC-BC, which are thought to involve a combination of transglycosylation and shifted binding with hydrolysis (Zhu et al., 1994). How the presence of domain C promotes the multimolecular hydrolytic reactions remains unknown, but it is possible that it sterically hinders the release of products from the initial hydrolytic event, thereby providing the opportunity for a transglycosylation reaction to ensue. If this were the case, then removal of domain C would expose the active site cleft and facilitate the more rapid release of hydrolysis products.
XynC-B was found to function optimally under conditions which are typical for many xylanases and to be expected within the rumen environment (39°C, pH 6.5) (Hobson, 1976). The data for temperature and pH optima obtained with XynC-B did not deviate significantly from the published values for the full XynC and XynC-BC (Zhu et al., 1994
), suggesting that, unlike the situation with the P.fluorescens enzymes (Black et al., 1996
), domain C does not confer stability to the catalytic domain. The subtle differences noted probably reflect the slightly different techniques used by the different researchers.
With the possibility of domain C contributing to stability being excluded, its role in XynC remains unknown. Presumably this domain endows the enzyme with a desirable feature because its presence nevertheless appears to be detrimental to the catalytic properties of domain B. An alternative role for this domain could involve proteinprotein interactions as provided by the dockerin domains present in other cellulolytic and xylanolytic enzymes. This postulate is based on previous observations indicating that the F.succinogenes enzymes form complexes on the cell surface (Paradis et al., 1993) and is supported in this study by HCA. While lacking any significant amino acid sequence homology and a hypothetical calcium binding motif (F-hand), the HCA plot of domain C is very similar to that of the dockerin domains of xylanase B and endoglucanase A produced by another ruminant bacterium, R.flavefaciens. The latter domains represent members of the type I dockerin domains found in the cellulolytic and xylanolytic enzymes of R.flavifaciens and species of Clostridium (Kirby et al., 1997
; Bayer et al., 1998
). Attractive as this postulate is, however, further detailed investigations are required to confirm the function of domain C of XynC as dockerin-like. The implication of such studies would be significant because XynC-C would provide the first example of a new class of dockerin-like domains.
The significance of linking multiple catalytic domains is not clear. In some instances, it may increase the efficiency of xylan hydrolysis to xylose by sequential activity of the domains, to generate oligosaccharides which function as substrates for the other. Clearly this does not seem to be the case for the F.succinogenes xylanase as the catalytic efficiency of the complete XynC is only a fraction of that of either XynC-A or XynC-B alone (Table III). Nevertheless, it is important to remember that these data were obtained from in vitro experiments in isolation from any potential factors, including the membrane environment of the cell surface, which may influence the activity of the enzyme and/or the individual domains. It has also been suggested that a multi-domain enzyme could confer a competitive advantage in that it may minimize release of hydrolysis products to competing microbes or to generate hydrolysis products that would not be generated as easily by one domain (Zhang and Flint, 1992
).
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Notes |
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2 Present address: Infectious Diseases Research, Lilly Research Laboratories, Lilly Corporate Center, Indianapolis, IN 46285, USA
3 To whom correspondence should be addressed. E-mail: aclarke{at}micro.uoguelph.ca
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Acknowledgments |
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References |
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Black,G.W., Rixon,J.E., Clarke,J.H., Hazlewood,G.P., Theodorou,M.K., Morris,P. and Gilbert,H.J. (1996) Biochem. J., 319, 515520.[ISI][Medline]
Bollag,D. and Edelstein,S. (1991) Protein Methods. Wiley, Toronto, pp. 116123.
Bray,M.R. and Clarke,A.J. (1994) Eur. J. Biochem., 219, 821827.[Abstract]
Clarke,A.J., Sarabia,V., Keenleyside,W., McLachlan,P.R. and Whitfield,C. (1991) Anal. Biochem., 199, 6874.[ISI][Medline]
Clarke,A.J. (1997) Biodegradation of Cellulose. Technomic, Lancaster, PA.
Gaboriaud,C., Bissery,V., Benchetrit,T. and Mornon,J.P. (1987) FEBS Lett., 224, 149155.[ISI][Medline]
Gilbert,H.J., Hazlewood,G.P., Laurie,J.I., Orpin,C.G. and Xue,G.P. (1992) Mol. Microbiol., 6, 20652072.[ISI][Medline]
Gilkes,N.R., Henrissat,B., Kilburn,D.G., Miller,R.C.,Jr and Warren,R.A.J. (1991) Microbiol. Rev., 55, 303315.[ISI]
Gong,J. and Forsberg,C. (1993) J. Bacteriol., 175, 68106821.[Abstract]
Guermeur,Y., Geourjon,C., Gallinari,P. and Deleage,G. (1999) Bioinformatics, 15, 413421.
Henrissat,B. (1999) Swiss-Prot Protein Sequence Data Bank, Release 38.0 and Updates to July 1999.
Henrissat,B. and Bairoch,A. (1993) Biochem. J., 293, 781788.[ISI][Medline]
Hill,T.W. (1996) Can. J. Microbiol., 42, 557561.[ISI]
Hobson,P.N. (1976) The Microflora of the Rumen. Meadowfield Press, London, pp. 115.
Kirby,J., Martin,J.C., Daniel,A.S. and Flint,H.J. (1997) FEMS Microbiol. Lett., 149, 213219.[ISI][Medline]
Laemmli,U.K. (1970) Nature, 227, 680685.[ISI][Medline]
Lemesle-Varloot,L., Henrissat,B., Gaboriaud,C., Bissery,V., Morgat,A. and Mornon,J.P. (1990) Biochemie, 72, 555574.[ISI][Medline]
McGavin,M.J. and Forsberg,C. (1988) J. Bacteriol., 170, 29142922.[ISI][Medline]
Nelson,N. (1994) J. Biol. Chem., 153, 375380.
Pagès,S., Belaich,A., Belaich,J.-P., Morag,E., Lamed,R., Shoham,Y. and Bayer,E.A. (1997) Proteins, 29, 517527.[ISI][Medline]
Paradis,F.W., Zhu,H., Krell,P.J., Phillips,J.P. and Forsberg,C.F. (1993) J. Bacteriol., 175, 76667672.[Abstract]
Somogyi,M. (1952) J. Biol. Chem., 195, 1923.
Xue,G.P., Gobius,K.S. and Orpin,C.G. (1992) J. Gen. Microbiol., 138, 23972403.[ISI][Medline]
Zhang,J.X. and Flint,H.J. (1992) Mol. Microbiol., 6, 10131023.[ISI][Medline]
Zhu,H., Paradis,F.W., Krell,P.J., Phillips,J.P. and Forsberg C.W. (1994) J. Bacteriol., 176, 38853894.[Abstract]
Received August 15, 1999; revised May 8, 2000; accepted June 10, 2000.