Module shuffling of a family F/10 xylanase: replacement of modules M4 and M5 of the FXYN of Streptomyces olivaceoviridis E-86 with those of the Cex of Cellulomonas fimi

Satoshi Kaneko1,2, Shinnosuke Iwamatsu3, Atsushi Kuno3,4, Zui Fujimoto5, Yoko Sato6, Kei Yura6, Mitiko Go6, Hiroshi Mizuno5, Kazunari Taira4,7, Tsunemi Hasegawa3, Isao Kusakabe7 and Kiyoshi Hayashi1

1 National Food Research Institute, Ministry of Agriculture, Forestry and Fisheries, 2–1–2 Kannondai, Tsukuba, Ibaraki 305-8642, 3 Department of Material and Biological Chemistry, Faculty of Science, Yamagata University, Yamagata 990-8560, 4 National Institute for Advanced Interdisciplinary Research, Ministry of International Trade and Industrial Science, Tsukuba, Ibaraki 305-8562, 5 National Institute of Agrobiological Resources, Ministry of Agriculture, Forestry and Fisheries, 2–1–2 Kannondai, Tsukuba, Ibaraki 305-8602, 6 Division of Biological Science, Graduate School of Science, Nagoya University, Furo-cho, Chikusa-ku, Nagoya, 464-8602 and 7 Institute of Applied Biochemistry, University of Tsukuba, 1–1–1 Tennoodai, Tsukuba, Ibaraki 305-8572, Japan


    Abstract
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To facilitate an understanding of structure–function relationships, chimeric xylanases were constructed by module shuffling between the catalytic domains of the FXYN from Streptomyces olivaceoviridis E-86 and the Cex from Cellulomonas fimi. In the family F/10 xylanases, the modules M4 and M5 relate to substrate binding so that modules M4 and M5 of the FXYN were replaced with those of the Cex and the chimeric enzymes denoted FCF-C4, FCF-C5 and FCF-C4,5 were constructed. The kcat value of FCF-C5 for p-nitrophenyl-ß-D-cellobioside was similar to that of the FXYN (2.2 s–1); however, the kcat value of FCF-C4 for p-nitrophenyl-ß-D-cellobioside was significantly higher (7.0 s–1). The loss of the hydrogen bond between E46 and S22 or the presence of the I49W mutation would be expected to change the position of Q88, which plays a pivotal role in discriminating between glucose and xylose, resulting in the increased kcat value observed for FCF-C4 acting on p-nitrophenyl-ß-D-cellobioside since module M4 directly interacts with Q88. To investigate the synergistic effects of the different modules, module M10 of the FCF-C4 chimera was replaced with that of the Cex. The effects of replacement of module M4 and M10 were almost additive with regard to the Km and kcat values.

Keywords: Cellulomonas fimi/Cex,chimeric xylanase/family 10 xylanase/module/Streptomyces olivaceoviridis


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Xylan is a major component of hemicelluloses in plant cell walls. It consists of a linear backbone of ß-1,4-linked xylopyranose units and has side chains that are often composed of other sugar residues such as arabinose and glucuronic acid. ß-Xylanase (EC 3.2.1.8) hydrolyses the ß-1,4-glycosidic linkages within the xylan backbone in an endo fashion. On the basis of the amino acid sequences of their catalytic domains, ß-xylanases have been classified into two glycanase families (F/10 and G/11) (Gilkes et al., 1991Go; Henrissat and Bairoch, 1993Go). So far, the three-dimensional structures of seven kinds of family F/10 xylanases have been solved (Derewenda et al., 1994Go; Harris et al., 1994Go; White et al., 1994Go; Dominguez et al., 1995Go; Schmidt et al., 1998Go; Natesh et al., 1999Go; Fujimoto et al., 2000Go). Family F/10 xylanases are known to have a (ß/{alpha})8-barrel structure and there are additional helices and loops which are arranged in a basic TIM barrel structure forming the active site cleft (Schmidt et al., 1998Go). Cellulomonas fimi endo-xylanase (Cex) is one of the most characterized xylanases (Tull et al., 1991Go; Bedarkar et al., 1992Go; MacLeod et al., 1994Go; White et al., 1994Go, 1996Go; MacLeod et al., 1996Go; Notenboom et al., 1998aGo, bGo). Based on the three-dimensional structure of Cex, the catalytic domain was divided into 22 modules (Sato et al., 1999Go). A module is a contiguous polypeptide segment of a protein which has a compact conformation within a globular domain. A module is defined by the distance between C{alpha} atoms and on average a module is about 15 amino acid residues long (Go, 1981Go; Go and Nosaka, 1987Go). Thirty-one intron sites of the family F/10 xylanase genes collected from fungi were found to correlate with the module boundaries with considerable statistical significance (Sato et al., 1999Go). This indicates that the location of the introns existing in eukaryotic xylanase genes are not random and supports the concept that introns play an important role in protein evolution as mediators of exon shuffling (Gilbert, 1978Go). Therefore, module shuffling in vitro mimics one of the natural mechanisms of protein evolution. Thus, we chose module shuffling as a tool in the elucidation of structure–function relationships of xylanases.

The enzyme FXYN from Streptomyces olivaceoviridis E-86 was selected as one of the parent enzymes since we have been successful in crystallizing the intact FXYN (Fujimoto et al., 1997Go, 2000Go) and also in isolating its gene (Kuno et al., 1998Go). In addition, the substrate specificity of the FXYN has been well characterized (Kusakabe et al., 1977Go, 1983Go; Yoshida et al., 1990Go, 1994Go; Matsuo et al., 1991Go). In a previous paper (Kaneko et al., 1999Go), we reported the effect of the replacement of module M10 because substrate binding amino acid residues had been identified by co-crystallization of Cex with an inhibitor (White et al., 1996Go) as being located in modules M4, M6, M7, M10, M15, M19 and M20 (Sato et al., 1999Go). In the present investigation, we selected modules M4 and M5 for shuffling. Modules M4 and M5 are one of the additional helices of the basic TIM barrel and are arranged in a loop to form part of the substrate-binding cleft of the enzyme (Figure 1Go). It is an important module for family F/10 xylanases because it contains amino acids which form the –1 to –3 sites of the substrate-binding cleft (Figure 2Go). To endeavour to understand the function of these modules, we constructed chimeric xylanases. In this paper, the function of modules M4 and M5 and the correlation of these results with those from module M10 are described.



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Fig. 1. Structures of the catalytic domains of FXYN and Cex and the position of modules M4 (red), M5 (pink) and M10 (blue) in the structures. (A) FXYN; (B) Cex. Glu128 and Glu236 in FXYN and Glu127 and Glu233 in Cex are the catalytic amino acids of the double displacement mechanism.

 


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Fig. 2. The –1 to –3 subsite in the substrate-binding cleft of FXYN. Hydrogen bonds between xylobiose and interacting amino acids are shown as dashed lines. Regions of modules M4 and M5 are indicated in brown.

 

    Materials and methods
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Construction of chimeric enzymes

The module arrangement of chimeric xylanases is shown in Figure 3Go. The catalytic domains of the FXYN and Cex xylanases were separately subcloned into the pQE60 vector (Qiagen, Hilden, Germany). Construction of the chimeras was performed by the polymerase chain reaction (PCR) using overlapping primers at their respective module boundaries (Kaneko et al., 1999Go). Briefly, DNA fragments from the FXYN coding modules M1 to M3 or M1 to M4 and modules M5 to M22 or M6 to M22 were amplified by PCR (Takara LA Taq, Takara Shuzo, Shiga, Japan). Each of the 25 amplification cycles consisted of denaturation at 98°C for 1 min and annealing and primer extension at 72°C for 1 min. DNA coding for module M4, module M5 or modules M4 and M5 of the Cex gene, with an overlapping region for the FXYN gene, were synthesized. The 10 bp overlapping regions of the primers were designed to be complementary at their respective module boundaries. The first round of PCR products were separated by agarose gel electrophoresis, followed by gel extraction, and used for the second round PCR without primers. Each of the 20 amplification cycles consisted of denaturation at 98°C for 1 min, annealing at 60°C for 25 min and primer extension at 72°C for 5 min. The strands having matching sequences at their respective module boundaries overlapped and acted as primers for each other. On the third round of PCR, the combined fragment was amplified by PCR primers with 25 cycles of shuttle PCR with denaturation at 98°C for 1 min and annealing and primer extension at 72°C for 1 min. The PCR products were subcloned into the plasmid pCR2.1 using the Original TA Cloning Kit and INV{alpha}F' cells (Invirogen, Carlsbad, CA), then the insert was sequenced with an automated DNA sequencer (Model 310, Applied Biosystems, Foster City, CA).



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Fig. 3. Construction of the chimeric xylanases and alignment of modules M4 and M5. Based on the three-dimensional structure of Cex, the catalytic domain of Cex was divided into 22 modules (Sato et al., 1999Go). The positions of the catalytic amino acid residues are depicted as {blacktriangledown}.

 
Production of enzymes in Escherichia coli

For expression in E.coli and purification of the FXYN, Cex, FCF-C4, FCF-C5, FCF-C4,5 and FCFCF-C4,10, the pET expression system (Novagen, Madison, WI) was employed. Each gene was individually inserted into the pET28a vector and the enzyme was expressed as a fusion protein consisting of the enzyme with a carboxyl-terminal tag with six histidine residues attached. Plasmids were used to transform E.coli BL21 (DE3) and the transformants were cultivated at 25°C in LB medium (1 l) containing kanamycin (20 µg/ml) until the optical density reached 0.4 at 600 nm. After the addition of isopropyl-1-thio-ß-D-galactoside (IPTG) to give a final concentration of 1 mM, the culture was incubated at 25°C for 24 h. The expressed enzymes were purified with a HisTrap chelating column (Pharmacia, Uppsala, Sweden). The enzyme was eluted from the column as a homogeneous protein by SDS–PAGE and the relevant fractions were pooled and dialysed against deionized water. The recovery of the FXYN, Cex, FCF-C4, FCF-C5 and FCFCF-C4,10 from 1 l of culture was 4.1, 0.1, 13.0, 0.04 4.4 mg, respectively.

Circular dichroism and steady-state kinetic studies

The circular dichroism spectra of the FXYN, Cex, FCF-C4, FCF-C5 and FCFCF-C4,10 enzymes were acquired using the conditions reported previously (Kaneko et al., 1999Go; Kuno et al., 1999Go). Steady-state kinetics were investigated as reported previously (Kaneko et al., 1999Go; Kuno et al., 1999Go). Briefly, the reaction mixture containing the substrate at various concentrations in 25% McIlvaine buffer (a mixture of 0.1 M citric acid and 0.2 M Na2HPO4, pH 7.0), containing 0.05% bovine serum albumin (BSA), was incubated at 30°C for 5 min before 50 µl of enzyme solution were added. The amount of p-nitrophenol released was determined by monitoring the absorbance at 400 nm with a spectrophotometer (DU-7400; Beckman, Palo Alto, CA). p-Nitrophenyl-ß-D-xylobioside (PNP-X2) was synthesized by the method described in a previous paper (Takeo et al., 1995Go). The xylobiose used in the synthesis was purified from `Xylobiose Mixture' (Suntory, Osaka, Japan). p-Nitrophenyl-ß-D-cellobioside (PNP-G2) was kindly donated by Yaizu Suisan (Yaizu, Japan).

Enzymatic properties

The effects of pH on the activity and stability of the various xylanases were investigated using a series of McIlvaine buffers (0.2 M Na2HPO4–0.1 M citric acid) from pH 4.0 to 8.0 and Atkins–Pantin buffers (0.2 M boric acid + 0.2 M KCl–0.2 M Na2CO3) from pH 8.0 to 10.5. The activities of the xylanases were assayed as follows. Each assay mixture contained 0.5 ml of a 2 mM PNP-X2 solution, 0.4 ml of buffer and 0.1 ml of enzyme solution. The reactions were performed at 45°C for 10 min whereupon they were stopped by the addition of 1.0 ml of 0.2 M Na2CO3 solution and the amount of p-nitrophenol (PNP) released was then determined at 408 nm. For determinations of the pH stabilities of the xylanases, the enzymes were pre-incubated in the absence of substrate at 30°C for 60 min and the residual activity was assayed at 45°C and pH 5.7. The effect of temperature on the activity of the xylanases was examined using a series of water-baths ranging from 30 to 70°C. Xylanase activity was measured at various temperatures at pH 5.7. For temperature-stability measurements of the various xylanases, the enzymes were pre-incubated at various temperatures at pH 7.0 for 60 min and the residual activity was then determined at 45°C and pH 5.7.

Xylan hydrolysis by chimeric xylanases

A reaction mixture containing 150 µl of McIlvaine buffer (pH 7.0), 50 µl of 1% (v/w) BSA and 250 µl of 1% soluble birchwood xylan solution was equilibrated at 30°C for 5 min and reactions were initiated by the addition of 50 µl of the enzyme (the final concentrations of FXYN, Cex, FCF-C4, FCF-C5 and FCF-C4, 10 were 0.012, 0.035, 0.023, 0.025 and 0.042 mg/ml, respectively. The amount of enzyme was adjusted by the production of reducing sugar from soluble birchwood xylan. After 0, 0.25, 0.5, 0.75, 1, 1.5, 3, 6, 12 and 24 h incubations, the reaction was terminated by boiling for 5 min. The reducing power generated from the soluble xylan was determined by the Somogyi–Nelson method (Somogyi, 1952Go). The reaction products were also analyzed using an HPAEC–PAD system (Dionex International Subsidiaries, Osaka, Japan).


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Construction and characterization of FCF-C4, FCF-C5 and FCF-C4,5

The overall similarity of the catalytic domains between the FXYN and Cex enzymes is 49%. As shown in Figure 1Go, the positions of modules M4 and M5 in the catalytic domain are in close proximity to catalytic amino acids such as E128 and E236 of the FXYN or to E127 and E233 of the Cex. Some of the amino acid residues forming the binding subsite are included in modules M4 and M5. The importance of these amino acids in substrate binding has been investigated by site-directed mutagenesis (Charnock et al., 1997Go, 1998Go). Therefore, we selected modules M4 and M5 for a shuffling investigation and chimeras FCF-C4, FCF-C5 and FCF-C4,5 were constructed (Figure 3Go). The activities of the constructed chimeric enzymes were detected on Remazole Brilliant Blue–xylan containing LB agar plates (data not shown). However, FCF-C4,5 was purified as inactive protein. The loss of activity of the enzyme during the purification step resulted from the high instability of the enzyme. Therefore, only FCF-C4 and FCF-C5 were purified and subjected to further investigations. Circular dichroism spectra obtained from the enzymes indicated that the constructed chimeric enzymes folded in the same conformational manner as the parent enzymes (Figure 4Go). The enzymatic properties of the FCF-C4 and FCF-C5 chimeras are shown in Figure 5Go. The optimum pHs of both chimeric enzymes were shifted to a slightly more acidic pH (from pH 6 of parental enzymes to pH 5), but the pH stability of the enzymes remained unchanged (Figure 5A and CGo). The optimum temperature of both FCF-C4 and FCF-C5 was 55°C, which was slightly lower than the 60°C observed for the parental FXYN and Cex enzymes (Figure 5BGo) and the thermostabilities of the chimeric enzymes were also slightly decreased compared with the parental enzymes (Figure 5DGo). Kinetic data were collected for both FCF-C4 and FCF-C5 along with parental FXYN and Cex using PNP-G2 and PNP-X2 as the substrates (Table IGo). Compared with the Cex, the FXYN enzyme displays much lower levels of activity towards both of the substrates. The FXYN enzyme displays kcat/Km values ~70-fold (for PNP-G2) and ~30-fold (for PNP-X2) lower than the corresponding Cex values. Because the majority of the chimeric enzymes FCF-C4 and FCF-C5 originated from the FXYN enzyme, the kinetic parameters of these enzymes would be expected to be closer to those of the parental FXYN rather than those of the Cex. The Km values of FCF-C4 and FCF-C5 for PNP-G2 were 112 and 96 mM, respectively, almost the same as that of the FXYN (97 mM). The kcat value of FCF-C5 for PNP-G2 (1.7 s–1) was also similar to the value of FXYN (2.2 s–1); however, surprisingly, the kcat value of FCF-C4 for PNP-G2 was significantly higher (7.0 s–1). The Km and kcat values of FCF-C4 and FCF-C5 for PNP-X2 were 0.88 mM and 21 s–1 and 2.4 mM and 33 s–1, respectively. The kcat/Km value of FCF-C4 was unchanged relative to FXYN. However, the kcat/Km value of FCF-C5 was slightly lower than that of FXYN, primarily owing to a reduction in the kcat value. The activities of the chimeric xylanases for the native substrates were also determined (Figure 6Go). When twice the amounts of FCF-C4 and FCF-C5 were used relative to FXYN, the enzyme activities were almost the same, indicating that the kcat values of the chimeric xylanases were almost half those of the FXYN enzyme. However, the mode of action of xylan hydrolysis by the chimeric enzymes was not altered, as evidenced by the analysis of the hydrolysis products by HPEAC–PAD (data not shown).



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Fig. 4. Circular dichroism spectra of the chimeric xylanases.

 


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Fig. 5. Enzymatic properties of the chimeric xylanases. (A) Effects of pH on enzyme activity; (B) effects of temperature on enzyme activity; (C) effects of pH on enzyme stability; (D) effects of temperature on enzyme stability. {circ}, Cex; {blacksquare}, FCF-C4; {blacktriangleup}, FCF-C5; •, FCFCF-C4,10; {triangleup}, FXYN.

 

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Table I. Kinetic parameters of chimeric xylanases
 


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Fig. 6. Xylan hydrolysis by the chimeric enzymes. {circ}, Cex; {blacksquare}, FCF-C4; {blacktriangleup}, FCF-C5; •, FCFCF-C4,10; {triangleup}, FXYN. Soluble birchwood xylan (10 mg/ml) was incubated at 30°C with enzyme concentrations of 0.012, 0.035, 0.023, 0.025 and 0.042 mg/ml for FXYN, Cex, FCF-C4, FCF-C5 and FCF-C4,10, respectively. After 0, 0.25, 0.5, 0.75, 1, 1.5, 3, 6, 12 and 24 h incubations, the reducing power generated from the soluble xylan was determined by the Somogyi–Nelson method (Somogyi, 1952Go).

 
Construction and characterization of FCFCF-C4,10

We previously reported the characterization of the chimera FCF-C10 in which module M10 of the FXYN was replaced with that of the Cex enzyme (Kaneko et al., 1999Go). Both the kcat and Km values observed for FCF-C10 were decreased by a factor of 10 relative to FXYN (Kaneko et al., 1999Go). To investigate the synergistic effects of the modules, module M10 of FCF-C4 was replaced with that of the Cex. The constructed chimera, FCFCF-C4,10, was expressed in an active form. FCFCF-C4,10 was purified and subjected to the same investigations as FCF-C4 and FCF-C5. Circular dichroism spectra of FCFCF-C4,10 indicated that the enzyme folded in the same manner as the parent enzymes (Figure 4Go). The enzymatic properties of the FCFCF-C4,10 chimera are depicted in Figure 5Go. The optimum pH of the chimeric enzyme was almost the same as that of the parental enzymes, but a difference in the pH stabilities of FCFCF-C4,10 and the FXYN was observed (Figure 5A and CGo). The stability of FCFCF-C4,10 in the mid-range of pH values was slightly lower in comparison with the other enzymes. Both the optimum temperature and the thermostability range of the FCFCF-C4,10 chimera were slightly reduced relative to the parental enzymes but were similar to those of the FCF-C4 and FCF-C5 enzymes (Figure 5B and DGo). The kinetic data obtained for the FCFCF-C4,10 chimera toward PNP-G2 and PNP-X2 are shown in Table IGo. Compared with the FCF-C4 chimera, FCFCF-C4,10 displays a lower activity towards both substrates. FCFCF-C4,10 displays a kcat/Km value ~2.5-fold lower for PNP-G2 and ~2.8-fold lower for PNP-X2 relative to the FCF-C4 chimera. The kinetic parameters of the FCFCF-C4,10 enzyme were closer to those of FCF-C10 than to those of FCF-C4. The Km value for FCFCF-C4,10 toward PNP-G2 was 46 mM, which was closer to that of FCF-C10 (64 mM) than to that for FCF-C4 (112 mM). The kcat value of FCFCF-C4,10 toward PNP-G2 (1.2 s–1) was also similar to the value observed for FCF-C10 (1.8 s–1). The Km value of FCFCF-C4,10 toward PNP-X2 was 0.14 mM, close to that observed for FCF-C10 (0.24 mM) and 6.3-fold lower than that seen for FCF-C4 (0.88 mM). The kcat value obtained for FCFCF-C4,10 toward the substrate PNP-X2 (1.2 s–1) was also reduced to a level 17.5-fold lower than that seen with FCF-C4 (21 s–1). The activities of the FCFCF-C4,10 chimera toward the native substrates were also determined (Figure 6Go). The enzyme activities of FCFCF-C4,10 were almost the same as those of the other enzymes when using twice the amount of enzyme as FCF-C4, indicating that the kcat value of FCFCF-C4,10 was almost half that of FCF-C4. However, the mode of action of xylan hydrolysis by the chimeric enzymes was unchanged, as evidenced by the analysis of the hydrolysis products by HPEAC–PAD (data not shown).


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The environment of modules M4 and M5 of the FXYN and Cex enzymes are shown in Figure 7A and BGo, respectively. Modules M4 and M5 in both the FXYN and Cex enzymes consist of eight and 13 amino acid residues, respectively. The identical amino acids in modules M4 and M5 between the FXYN and Cex are 75% and 54%, respectively. As shown in Figure 2Go, module M4 of FXYN includes amino acids such as E44, N45 and K48, which are directly involved in substrate binding (White et al., 1996Go; Charnock et al., 1997Go, 1998Go). These amino acids are conserved in both the FXYN and Cex enzymes. When module M4 of FXYN was replaced with that of Cex, only two amino acids were replaced (E46A and I49W). These mutations did not significantly affect protein folding or the enzymatic properties of the enzyme (Figures 4 and 5GoGo). There being only two amino acids replaced, the kcat value of FCF-C4 for PNP-G2 was significantly increased relative to that of the parent FXYN (Table IGo). In FXYN, Q88 hydrogen bonds to D50 and K48 (Figure 7AGo). A hydrogen bond between E46 and S22 is also observed (Figure 7AGo). In contrast, in Cex the former hydrogen bonds are observed, but the latter is not possible in Cex since the corresponding amino acid to E46 in FXYN is A45 in Cex (Figure 7BGo). Further differences in the structures of the FXYN and Cex enzymes were also observed. Residue I49 in FXYN is close to M93 in space. I49 in FXYN corresponds to residue W48 in Cex and W48 possesses a larger side chain than I49; therefore, in order to avoid steric hindrance, the residue corresponding to M93 in FXYN is actually A92 in Cex, which has a small side chain. The I49W mutation observed in the FCF-C4 chimera would be expected to have an effect on the structure of module M4. Previously, it has been reported that the glutamine residue Q87 of Cex is a key residue in discriminating between glucose and xylose at the –2 subsite (Notenboom et al., 1998aGo). Very recently, the corresponding residue, namely Y87 and Q88 in Pseudomonas fluorescens XYLA and Streptomyces lividans XynA, respectively was also proved to play a pivotal role in discriminating between glucose and xylose (Andrews et al., 2000Go; Ducros et al., 2000Go). The loss of the hydrogen bond between residues E46 and S22 or the I49W mutation in FXYN would be expected to change the spatial position of Q88 in FXYN, resulting in the increase observed in the kcat value of FCF-C4 for PNP-G2 since module M4 interacts directly with Q88 in FXYN. However, the structural changes in the substrate-binding cleft did not affect the substrate specificity toward xylan. These results are in accordance with an earlier report which describes E43, N44 and K47 in XYLA of P.fluorescens which corresponding with E44, N45 and K48 in FXYN as having a critical role in the activity of P.fluorescens XYLA against xylooligosaccharides but not against highly polymeric substrates such as xylan (Charnock et al., 1997Go). In contrast, when module M5 was replaced, the kinetic parameters did not change significantly in spite of six amino acids being altered. There is hydrogen bonding between E53 and H108 in FXYN; the replaced amino acids did not affect the bonding. It seems to be that the replacement of the module M5 did not affect the structure of the enzyme.



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Fig. 7. The environment of modules M4 and M5 in FXYN and Cex. (A) FXYN; (B) Cex. Hydrogen bonds are indicated as dashed lines.

 
It has been reported that as multiple mutations were introduced to thermolysin, the logarithm of the activity was found to be almost additive (Kidokoro, 1998Go). Therefore, we tried to determine whether or not the module replacement is also additive. Module M10 of FCF-C4 was also replaced (FCFCF-C4,10) since we have previously reported the effect of the replacement of module M10 (Kaneko et al., 1999Go). As shown in Figure 1Go, the location of modules M4 and M10 was distinct and these modules did not interact with each other so that these modules are independent of each other. The enzymatic properties of the FCFCF-C4,10 chimera did not change significantly in relation to those of FXYN and FCF-C4. The enzyme stability of FCFCF-C4,10 at neutral pH was slightly decreased relative to the parental FXYN. The reason cannot be deduced only from the data in this paper, but it may be due to slight distortion caused by the double replacement of both modules M4 and M10. We determined that the Km and kcat values of FCF-C10 for PNP-X2 were reduced 10-fold relative to those of FXYN; however, kcat/Km did not change (Kaneko et al., 1999Go).

When the module M10 of FCF-C4 was replaced with that of Cex, the Km value for PNP-X2 was also decreased from 0.88 to 0.14 mM together with kcat value from 21 to 1.2 s–1. There was a similar situation when module M10 of FXYN was replaced with that of Cex. Therefore, the effect of module M10 was almost the same for FXYN and FCF-C4 so that in FCFCF-C4,10, the effects of replacement of module M4 and M10 were almost additive.

In conclusion, our results indicate that in xylanases, module M4, including the substrate-binding residues (E44, N45 and K48 in FXYN), is not only related directly to enzyme activity but also interacts with adjacent modules which are involved in forming the substrate-binding cleft. A similar finding was apparent in the case of module M10 replacement (Kaneko et al., 1999Go). The double replacement of modules M4 and M10 displayed additive effects in Km and kcat for replacing modules M4 and M10 individually. These results indicate that module shuffling or exon suffling is a possible mechanism in the evolution of proteins.


    Notes
 
2 To whom correspondence should be addressed. E-mail: sakaneko{at}nfri.affrc.go.jp Back


    Acknowledgments
 
The authors thank Suntory for the supply of `Xylobiose Mixture' and Dr Yasuo Gama, NIMC, AIST, for help with the synthesis of PNP-X2. We are also grateful for the editorial assistance of Dr Joanne B.Hart in the preparation of the manuscript. This work was supported in part by the Program for Promotion of Basic Research Activities for Innovative Biosciences.


    References
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
Andrews,S.R., Charnock,S.J., Lakey,J.H., Davies,G.J., Claeyssens,M., Nerinckx,W., Underwood,M., Sinnott,M.L., Warren,R.A.J. and Gilbert,H.J. (2000) J. Biol. Chem., 275, 23027–23033.[Abstract/Free Full Text]

Bedarkar,S., Gilkes,N.R., Kilburn,D.G., Kwan,E., Rose,D.R., Miller,R.C.,Jr, Warren,R.A. and Withers,S.G. (1992) J. Mol. Biol., 228, 693–695.[ISI][Medline]

Charnock,S.J., Lakey,J.H., Virden,R., Hughes,N., Sinnott,M.L., Hazlewood,G.P., Pickersgill,R. and Gilbert,H.J. (1997) J. Biol. Chem., 272, 2942–2951.[Abstract/Free Full Text]

Charnock,S.J., Spurway,T.D., Xie,H., Beylot,M.H., Virden,R., Warren,R.A., Hazlewood,G.P. and Gilbert,H.J. (1998) J. Biol. Chem., 273, 32187–32199.[Abstract/Free Full Text]

Derewenda,U., Swenson,L., Green,R., Wei,Y., Morosoli,R., Shareck,F., Kluepfel,D. and Derewenda,Z.S. (1994) J. Biol. Chem., 269, 20811–20814.[Abstract/Free Full Text]

Dominguez,R., Souchon,H., Spinelli,S., Dauter,Z., Wilson,K.S., Chauvaux,S., Beguin,P. and Alzari,P.M. (1995) Nature Struct. Biol., 2, 569–576.[ISI][Medline]

Ducros,V., Charnock,S.J., Derewenda,U., Derewenda,Z.S., Dauter,Z., Dupont,C., Shareck,F., Morosoli,R., Kluepfel,D. and Davies,G.J. (2000) J. Biol. Chem., 275, 23020–23026.[Abstract/Free Full Text]

Fujimoto,Z., Mizuno,H., Kuno,A., Yoshida,S., Kobayashi,H. and Kusakabe,I. (1997) J. Biochem. (Tokyo), 121, 826–828.[Abstract]

Fujimoto,Z., Kuno,A., Kaneko,S., Yoshida,S., Kobayashi,H., Kusakabe,I. and Mizuno,H. (2000) J. Mol. Biol., 300, 575–585.[ISI][Medline]

Gilbert,W. (1978) Nature, 271, 501.[ISI][Medline]

Gilkes,N.R., Henrissat,B., Kilburn,D.G., Miller,R.C.,Jr and Warren,R.A.J. (1991) Microbiol Rev., 55, 303–315.[ISI]

Go,M. (1981) Nature, 291, 90–92.[ISI][Medline]

Go,M. and Nosaka,M. (1987) Cold Spring Harbor Symp. Quant. Biol., 52, 915–924.[ISI][Medline]

Harris,G.W., Jenkins,J.A., Connerton,I., Cummings,N., Lo Leggio,L., Scott,M., Hazlewood,G.P., Laurie,J.I., Gilbert,H.J. and Pickersgill,R.W. (1994) Structure, 2, 1107–1116.[ISI][Medline]

Henrissat,B. and Bairoch,A. (1993) Biochem. J., 293, 781–788.[ISI][Medline]

Kaneko,S. et al. (1999) FEBS Lett., 460, 61–66.[ISI][Medline]

Kidokoro,S. (1998) Adv. Biophys., 35, 121–143.[ISI][Medline]

Kuno,A., Shimizu,D., Kaneko,S., Koyama,Y., Yoshida,S., Kobayashi,H., Hayashi,K., Taira,K., Kusakabe,I. (1998) J. Ferment. Bioeng., 86, 434–439.[ISI]

Kuno,A., Shimizu,D., Kaneko,S., Hasegawa,T., Gama,Y., Hayashi,K., Kusakabe,I. and Taira,K. (1999) FEBS Lett., 450, 299–305.[ISI][Medline]

Kusakabe,I., Kawaguchi,M., Yasui,T. and Kobayashi,T. (1977) Nippon Nogeikagaku Kaishi, 51, 429–437.

Kusakabe,I., Ohgushi,S., Yasui,T. and Kobayashi,T. (1983) Agric. Biol. Chem., 47, 2713–2723.[ISI]

MacLeod,A.M., Lindhorst,T., Withers,S.G. and Warren,R.A. (1994) Biochemistry, 33, 6371–6376.[ISI][Medline]

MacLeod,A.M., Tull,D., Rupitz,K., Warren,R.A. and Withers,S.G. (1996) Biochemistry, 35, 13165–13172.[ISI][Medline]

Matsuo,N., Yoshida,S., Kusakabe,I. and Murakami,K. (1991) Agric. Biol. Chem., 55, 2905–2907.[ISI][Medline]

Natesh,R., Bhanumoorthy,P., Vithayathil,P.J., Sekar,K., Ramakumar,S. and Viswamitra,M.A. (1999) J. Mol. Biol., 288, 999-1012.[ISI][Medline]

Notenboom,V., Birsan,C., Warren,R.A.J., Withers,S.G. and Rose,D.R. (1998a) Biochemistry, 37, 4751–4758.[ISI][Medline]

Notenboom,V., Brirsan,C., Nitz,M., Rose,D.R., Warren,R.A.J. and Withers,S.G. (1998b) Nature Struct. Biol., 5, 812–818.[ISI][Medline]

Sato,Y., Niimura,Y., Yura,K. and Go,M. (1999) Gene, 238, 93–101.[ISI][Medline]

Schmidt,A., Schlacher,A., Steiner,W., Schwab,H. and Kratky,C. (1998) Protein Sci., 7, 2081–2088.[Abstract/Free Full Text]

Somogyi,M. (1952) J. Biol. Chem., 195, 19–23.[Free Full Text]

Takeo,K., Ohguchi,Y., Hasegawa,R. and Kitamura,S. (1995) Carbohydr. Res., 277, 231–244.[ISI][Medline]

Tull,D., Withers,S.G., Gilkes,N.R., Kilburn,D.G., Warren,R.A. and Aebersold,R. (1991) J. Biol. Chem., 266, 15621–15625.[Abstract/Free Full Text]

White,A., Withers,S.G., Gilkes,N.R. and Rose,D.R. (1994) Biochemistry, 33, 12546–12552.[ISI][Medline]

White,A., Tull,D., Johns,K., Withers,S.G. and Rose,D.R. (1996) Nature Struct. Biol., 3, 149–154.[ISI][Medline]

Yoshida,S., Kusakabe,I., Matsuo,N., Shimizu,K., Yasui,T. and Murakami,K. (1990) Agric. Biol. Chem., 54, 449–457.[ISI][Medline]

Yoshida,S., Ono,T., Matsuo,N. and Kusakabe,I. (1994) Biosci. Biotechnol. Biochem., 58, 2068–2070.[ISI][Medline]

Received June 30, 2000; revised October 26, 2000; accepted October 30, 2000.





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