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Journal of Bacteriology, March 2009, p. 1641-1649, Vol. 191, No. 5
0021-9193/09/$08.00+0     doi:10.1128/JB.01285-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

The Membrane Lipoprotein LppX of Paenibacillus sp. Strain W-61 Serves as a Molecular Chaperone for Xylanase of Glycoside Hydrolase Family 11 during Secretion across the Cytoplasmic Membrane{triangledown} ,{dagger}

Mutsumi Fukuda,1,{ddagger} Seiji Watanabe,1,{ddagger},§ Jun Kaneko,1 Yoshifumi Itoh,1,{ddagger} and Yoshiyuki Kamio1,2*

Laboratory of Applied Microbiology, Department of Microbial Biotechnology, Graduate School of Agricultural Science, Tohoku University, 1-1 Tsutsumidori Amamiyamachi, Aobaku, Sendai 981-8555,1 Department of Human Health and Nutrition, Graduate School of Comprehensive Human Sciences, Shokei Gakuin University, Yurigaoka 4-10-1, Natori 981-1295, Japan2

Received 12 September 2008/ Accepted 9 December 2008


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ABSTRACT
 
Paenibacillus sp. strain W-61, which can utilize xylan as the sole source of carbon and energy, produces extracellular xylanases 1 and 3 (Xyn1 and Xyn3) and cell surface xylanase 5. In this study we found that lppX, immediately downstream of xyn1, encodes a lipoprotein located on the outer layer of the cytoplasmic membrane and that the LppX lipoprotein is essential for the secretion of active Xyn1 across the cytoplasmic membranes. In Escherichia coli, wild-type LppX was destined for the inner layer of the outer membrane. Mutant LppX(C19A), in which Cys-19, a possible lipomodification residue, is replaced with Ala, was located in the periplasm without being anchored to the membranes. Another mutant, LppX(S20D S21D), with substitutions of Asp for Ser-20 and Ser-21 (conversion to an Asp-Asp signal for sorting to the inner membrane), resided on the outer layer of the inner membrane, demonstrating that LppX has the sorting property of a lipoprotein. E. coli harboring both xyn1 and lppX secreted active Xyn1 into the periplasm. In contrast, E. coli carrying xyn1 alone failed to do so, accumulating inactive Xyn1 in the cytoplasmic membranes. Exogenous LppX(C19A) liberated the inactive Xyn1, which had been stagnating in the inner membrane, into the medium as an active enzyme. Thus, we propose that LppX is a novel type of lipoprotein that assists Xyn1 in making the proper fold necessary for traveling across the cytoplasmic membranes to be secreted as an active enzyme.


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INTRODUCTION
 
Some microorganisms produce several xylanolytic enzymes to hydrolyze β-1,4-xylan backbones and side chain sugars of hemicellulose in plant cell walls, so that constituent xyloses and other sugars can be utilized as carbon and energy sources (31). The xylan-degrading bacterium Paenibacillus sp. strain W-61 (formerly Aeromonas caviae W-61) produces five extracellular xylanases (Xyn1 to Xyn5) (25, 38-40, 45). Xyn1 and Xyn3 are secreted into the medium, where the latter is proteolytically converted into Xyn2 (40). Xyn5 is associated with the cell during the exponential-growth phase, but in the stationary phase it is released into the medium and is subsequently cleaved by proteolysis to form Xyn4 (25).

Xyn1 belongs to glycoside hydrolase (GH) family 11, a predominant group of small (<30-kDa) and basic xylanases of diverse origins, including fungi (such as Aspergillus, Penicillium, and Trichoderma spp.) as well as gram-positive (Bacillus, Clostridium, and Paenibacillus spp.) and gram-negative (Pseudomonas and Fibrobacter spp.) bacteria (10). In many organisms, the synthesis of xylanase is induced in the presence of the substrate xylan, probably via the products of its action, and it is subjected to catabolite repression by glucose (10). In Bacillus stearothermophilus no. 236, besides regulation by an inducer and a catabolite repressor at transcription, transcript levels of xynA (encoding a GH family 11 xylanase) are posttranscriptionally modulated by XaiF, the product of a gene immediately downstream of xynA (26). This protein prevents the decay of xynA mRNA caused by RNase and thereby increases XynA production (26).

Because of their potential applications in pulp, paper, food, and other industries (2), large quantities of fungal and bacterial xylanases have been generated in heterologous prokaryotic and eukaryotic hosts. Successful production of recombinant GH family 11 xylanases (2) indicates that additional factors, such as chaperone and secretion machines, are not required for their secretion by heterologous hosts (or by their native hosts).

In this study we determined the nucleotide sequence of the downstream gene and the cellular location of its product. A polypeptide encoded by the downstream gene had 61% amino acid identity to XaiF and was essential for Xyn1 production by strain W-61. The polypeptide was found to be a lipoprotein and to be located on the outer layer of the cytoplasmic membrane. We therefore designated this gene lppX (for "lipoprotein for Xyn1 production"). In accordance with its membrane location, LppX lipoprotein had no stabilizing activity toward xyn1 mRNA. We showed that LppX could be delivered to the cytoplasmic or outer membranes of Escherichia coli depending on the second and third amino acids of the sorting signal, demonstrating that LppX has the membrane-sorting property of a lipoprotein. Importantly, LppX did liberate inactive Xyn1, which had accumulated in the cytoplasmic membranes of an lppX mutant, into the medium as an active enzyme. These results led us to propose that lipoprotein LppX helps Xyn1 in traversing the cytoplasmic membrane as an active enzyme. To our knowledge, LppX is the first example of a lipoprotein with chaperone activity.


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MATERIALS AND METHODS
 
Bacterial strains, plasmids, and media. Table 1 lists the bacterial strains and plasmids used in the present study. Paenibacillus sp. strain W-61 and its derivatives were cultured in minimal medium (Medium I) (38) containing 0.7% (wt/vol) water-soluble xylan as a carbon source (or another specified carbon source) or in Luria-Bertani (LB) medium (46): Medium I was used in growth tests and for the preparation of the total RNA and cell proteins used in Northern and Western blotting, respectively, and LB medium was used for the preparation of chromosomal and plasmid DNAs and in transformation. Escherichia coli strains were routinely cultured in LB medium. Water-soluble xylan was prepared from oat-spelt xylan (Nacalai Tesque, Kyoto, Japan) as described previously (38). When appropriate, antibiotics were added to cultures at the indicated concentrations. Cultures were incubated aerobically at 37°C.


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  		TABLE 1. Bacterial strains and plasmids

Cloning and DNA sequencing of xyn1 flanking regions. The flanking regions of xyn1 on Paenibacillus sp. strain W-61 chromosomal DNA were cloned by inverse PCR walking as described in a previous paper (52). Nucleotide sequences of the amplified DNA fragments were determined using the ABI Prism BigDye Terminator cycle sequencing ready reaction kit and an ABI 377 DNA sequencer (Applied Biosystems, Foster City, CA). The sequences determined were assembled and analyzed using GENETYX-Mac ATSQ software and the GENETYX program (both from Genetyx Co., Tokyo, Japan), respectively. Homology searches were performed using the FASTA and BLAST programs implemented at the DDBJ/EMBL/GenBank or Swiss-Prot/NBRF-PIR databases.

Expression of xyn1 and xyn1-lppX in E. coli. The DNA regions of xyn1 and xyn1-lppX were amplified by PCR using Paenibacillus sp. strain W-61 chromosomal DNA as a template and primers pairs P1-P2 and P1-P3 (Table 2), respectively. After digestion at the BamHI and HindIII sites created by the primers, the PCR products were inserted into plasmid pUC119 between the corresponding sites to construct plasmids pX1T (xyn1) and pXFT (xyn1-lppX) (Table 1). Plasmid DNA was introduced into E. coli DH5{alpha} by transformation (6).


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  		TABLE 2. Oligonucleotide primers

Construction of an lppX knockout mutant. A 2.5-kbp DNA fragment covering xyn1 and lppX was amplified by PCR using Paenibacillus sp. strain W-61 chromosomal DNA as a template and primers P1 and P4 (Table 2). The resultant PCR products were doubly digested with BamHI and HindIII and were cloned between the same sites of plasmid pHY300PLK. Then an internal 740-bp StuI-and-BanII region of lppX was replaced with a cat gene fragment, which had been PCR amplified from plasmid pC194 (23) using primers P5 and P6 (Table 2): the protruding ends of both the insert and vector DNA fragments were made blunt by using T4 DNA polymerase (TaKaRa Bio, Otsu, Japan). The lppX::cat mutant gene thus created was recovered together with xyn1 on a BamHI-HindIII fragment from the plasmid and was cloned, after being converted to a blunt-end fragment as described above, into the temperature-sensitive shuttle vector pKAF at the SmaI site, to create plasmid pKMC (Table 1). This temperature-sensitive plasmid was introduced into Paenibacillus sp. strain W-61 by electroporation (6), and the transformed culture was cultivated in LB broth containing chloramphenicol (10 µg/ml) at 43°C to cure pKMC. The cured culture was spread onto LB plates containing chloramphenicol (10 µg/ml) to select strain PSC301 (lppX::cat), which had acquired the mutated gene via single crossover at xyn1, as verified by Southern blotting (25). Because xyn1 on pKMC lacks the promoter for xyn1-lppX, the chromosomal lppX is not expressed.

A plasmid carrying lppX fused to the xyn1 promoter was constructed for use in complementation tests. A putative promoter region of xyn1 (an intergenic region between orf4 and xyn1 and nucleotides –185 to –10 relative to the translation initiation codon) was amplified using Paenibacillus sp. strain W-61 chromosomal DNA as a template and primers P7 and P8 (Table 2). The amplified DNA fragments were cleaved with EcoRI and BamHI and were cloned into plasmid pHY300PLK between the corresponding sites. A DNA segment spanning the region from the ribosome-binding site to a putative transcription termination site (nucleotides –24 to +898) of lppX was PCR amplified using Paenibacillus sp. strain W-61 chromosomal DNA as a template and primers P9 and P4 (Table 2). The amplified DNA fragments were digested with BamHI and HindIII and were inserted between the BamHI site at the 3' end of the xyn1 promoter (Pxyn1) and the HindIII site in the polylinker of the vector, to produce plasmid pHPX4T (Pxyn1::lppX). Although Pxyn1 is a very weak promoter, when the Pxyn1::lppX fusion was cloned into the high-copy-number plasmid pHY300PLK, it was expressed at significantly high levels (see Results).

Construction of lppX mutants. Site-directed mutagenesis of lppX proceeded by the overlapping-extension method (20) using W-61 chromosomal DNA as a template and three pairs of primers, P10 and P11, P12 and P13, and P14 and P15, which introduce amino acid changes of Cys-19 to Ala, Ser-20 to Asp, and Ser-20 and Ser-21 to Asp (Table 2; the alanine and aspartate codons, respectively, are underlined). The mutated xyn1-lppX DNA fragments were then synthesized using primers P1 and P2 (Table 2). After digestion with BamHI-HindIII, the PCR products were inserted into plasmid pUC119 between the same sites. The resultant plasmids carrying a xyn1-lppX fragment with C19A or S20D S21D mutations in lppX were designated pC19A and pS20DS21D, respectively (Table 1). Likewise, the lppX(C19A) sequence was PCR amplified using primers P16 and P17 (Table 2), and after removal of the 3' protrusion created during PCR using T4 DNA polymerase and subsequent digestion with XhoI, the amplified fragments were cloned into plasmid pET-15b between the XhoI site and the blunted NcoI site, to produce plasmid pC19A2 (Table 2). Plasmids were transformed into E. coli BL21(DE3) as described previously (6).

Western and Northern blotting. An antiserum against LppX was raised in mice as described previously (25). Xyn1 and LppX, resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), were transferred to Hybond ECL membranes and cross-reacted with antisera against Xyn1 and LppX, respectively, as previously described (40). The signal intensities of protein bands on Western blots were quantified using NIH Image (http://rsbweb.nih.gov/nih-image/) (52). Total RNA was extracted by the methods described by Aiba et al. (1), and Northern blotting was performed using ECL Random Prime and Gene Image CDP-Star detection kits (GE Healthcare, Buckinghamshire, United Kingdom) according to the supplier's instructions.

Labeling of LppX with [14C]palmitic acid in vivo. [1-14C]palmitic acid (1.85 MBq; Daiichi Pure Chemicals, Tokyo, Japan) was added at a final concentration of 10 nM (185 kBq/ml) to early-log-phase cultures (optical density at 600 nm, 0.2) of strain PSC301/pHY300PLK or PSC301/pHPX4T in 100 ml of Medium I containing 0.7% (wt/vol) glucose and tetracycline (5 µg/ml). The cultures were incubated to reach the early-stationary phase (optical density at 600 nm, 2.5). Labeled cells were harvested by centrifugation, washed twice with 50 mM Tris-HCl buffer (pH 8.0) at 4°C, and homogenized on ice using a mortar and pestle in the presence of sea sand. After incubation with DNase I and RNase A (each at 50 µg/ml) at room temperature for 20 min, the cell lysates were centrifuged at 4°C and 1,000 x g for 5 min to remove the sea sand. The resultant supernatants were further centrifuged at 4°C and 200,000 x g for 1 h to separate cytosolic proteins and cytoplasmic membranes. The pelleted cytoplasmic membranes were washed with 50 mM Tris-HCl buffer (pH 8.0) and dissolved in 75 µl of 10% SDS. Cytosolic and membrane proteins were resolved by SDS-PAGE, and radioactive proteins in the gels were detected using a FIA-2000 fluoroimaging analyzer (Fujifilm, Tokyo, Japan).

Preparation of protoplasts and inside-out vesicles of cytoplasmic membranes from Paenibacillus sp. strain W-61. Protoplasts of PSC301/pHPX4T were prepared as described by Egelseer et al. (12). PSC301/pHPX4T cells growing exponentially in Medium I containing soluble xylan and tetracycline (5 µg/ml) were collected by centrifugation, suspended in 50 mM Tris-HCl (pH 8.0) containing 20 mM MgCl2 and 7.5% (wt/vol) polyethylene glycol 6000 (Wako Pure Chemical Industries, Osaka, Japan) (buffer A), and then incubated at 37°C for 60 min in the presence of egg white lysozyme (40 µg/ml). Protoplast formation was monitored by phase-contrast microscopy; almost all cells became protoplasts within 60 min. The protoplasts were harvested by centrifugation at 7,000 x g for 10 min at room temperature and were suspended in buffer A. To prepare inside-out vesicles of the cytoplasmic membranes, the protoplasts were suspended in 5 ml of 50 mM Tris-HCl (pH 8.0) containing 20 mM MgCl2 (buffer B) and were passed through a French pressure cell at 13,000 lb/in2, as described by Futai (17). The resultant inside-out membrane vesicles were collected by centrifugation at 4°C and 100,000 x g for 120 min and were dissolved in 4 ml of buffer B.

Preparation of periplasmic proteins, spheroplasts, and outer and inner membrane vesicles of E. coli. Spheroplasts were prepared as described by Osborn et al. (41) from exponentially growing E. coli cells (5 x 108/ml) in 1 liter of LB broth containing 0.5% (wt/vol) glucose and 100 µg/ml ampicillin at 37°C. The cells were suspended in 14 ml of ice-cold 10 mM Tris-HCl buffer (pH 7.5) containing 750 mM sucrose and lysozyme (100 µg/ml) and were then placed on ice for 2 min. Thereafter, 28 ml of 1.5 mM ice-cold EDTA (pH 7.5) was added to the cell suspension at a rate of 2.8 ml/min. After 30 min on ice, nearly all cells became spheroplasts as determined by microscopy. The spheroplasts were then separated from periplasmic proteins and outer membranes by centrifugation at 4°C and 8,000 x g for 10 min. Pelleted spheroplasts were suspended in 10 mM Tris-HCl buffer (pH 7.5) containing 250 mM sucrose and were disrupted by sonication to prepare inside-out vesicles of the inner membranes (17). The membrane vesicles were collected by ultracentrifugation at 200,000 x g for 60 min.

The inside-out vesicles of the outer membranes in the supernatant were pelleted by centrifugation again at 4°C and 200,000 x g for 60 min, and periplasmic proteins were obtained from the supernatant. Inside-out vesicles of the outer and the inner membranes were also prepared from exponentially proliferating E. coli cells as described above using sucrose density gradient centrifugation (35). The amounts of a 33-kDa major outer membrane protein and cytochrome b1 in the inner and the outer membrane vesicles were <0.1% (wt/vol) of those in the other preparation.

Proteinase K digestion of intact cells and membranes of Paenibacillus sp. strain W-61 and E. coli. Intact cells and membrane vesicles were digested with proteinase K at appropriate concentrations (see legends to Fig. 3, 5, and 6). At various times during the incubation period, 100-µl portions of reaction mixtures were transferred to Eppendorf tubes containing 10 µl of 40 mM phenylmethylsulfonyl fluoride to terminate the reaction. The amounts of Xyn1 and LppX in the cells and the membrane vesicles were determined by Western blotting as described above.


Figure 3
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  		FIG. 3. Location of LppX on the outer layer of the cytoplasmic membrane of Paenibacillus sp. strain W-61. Protoplasts and inside-out vesicles of the cytoplasmic membranes were prepared from W-61 cells growing exponentially in Medium I with water-soluble xylan. (A) The protoplasts were incubated at 20°C with or without 12.5 µg proteinase K in 4 ml of 50 mM Tris-HCl (pH 8.0) containing 20 mM MgCl2 and 7.5% (wt/vol) polyethylene glycol. (B) The inside-out membrane vesicles were treated with proteinase K as described in the legend to panel A. One reaction mixture also contained 1% (wt/vol) SDS to solubilize the membrane vesicles. At the indicated times, 100-µl portions of reaction mixtures were withdrawn and mixed with phenylmethylsulfonyl fluoride (final concentration, 4 mM) to terminate proteinase digestion. The amounts of LppX that remained undigested were determined by Western blotting.


Figure 5
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  		FIG. 5. Cellular locations of LppX and its mutant proteins in E. coli. (A) Soluble proteins in the cytoplasm and periplasm (S), inner membranes (IM), and outer membranes (OM) were prepared from 5 x 108 E. coli DH5{alpha}/pXFT (xyn1-lppX) cells and analyzed for the presence of LppX by Western blotting. (B) Intact cells (IC) (5 x 108 cells) and inside-out outer membrane vesicles (OV) (0.5 mg of protein) of E. coli DH5{alpha}/pXFT were incubated with or without proteinase K (50 µg/ml) in 4 ml of buffer B (see Materials and Methods) at 20°C for 10 min. Portions (10 µl) of reaction mixtures were analyzed for LppX by Western blotting. (C and D) The amounts of LppX in the periplasm (P) and the inner (IM) and outer (OM) membranes of E. coli DH5{alpha} (5 x 108 cells) harboring pC19A (C) or pS20DS21D (D) were analyzed by Western blotting.


Figure 6
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  		FIG. 6. Cellular locations of Xyn1 synthesized by E. coli in the presence or absence of lppX. (A) Spheroplasts (S) and periplasmic proteins (P) were prepared from late-exponential-growth-phase cells of E. coli DH5{alpha}/pX1T (xyn1) and DH5{alpha}/pXFT (xyn1-lppX). The spheroplasts and periplasmic proteins from 5 x 106 cells of each strain were analyzed by Western blotting for their Xyn1 contents. (B) The spheroplasts and inside-out cytoplasmic membrane vesicles of E. coli DH5{alpha}/pX1T cells (5 x 108 cells) were incubated with proteinase K (50 µg/ml) in 4 ml of buffer B containing 7.5% (wt/vol) polyethylene glycol at 20°C for 10 min, and amounts of Xyn1 that had escaped digestion by the proteinase were determined by Western blotting. The molecular sizes of Xyn1 (21 kDa) and its degradation product (19 kDa) were determined from their mobilities relative to those of molecular markers.

Activation of Xyn1 by LppX in vitro. We adopted the conditions for assay of the p20 (LolB) activity in releasing LppX lipoprotein on spheroplasts (34) to examine the chaperone activity of LppX for inactive Xyn1 on E. coli spheroplasts. Reaction mixtures (0.85 ml) contained 5 x 108 E. coli DH5{alpha}/pX1T spheroplasts, 0.5 mg periplasmic proteins of E. coli BL21(DE3)/pC19A, 62.5 mM Tris-HCl (pH 7.0), and 156 mM sucrose. After incubation at 30°C for 10 min, the reaction mixtures were chilled on ice and then centrifuged at 4°C and 20,000 x g for 10 min to separate spheroplasts in the pellets and soluble proteins in the supernatant. The amount and activity of Xyn1 protein were determined by Western blotting using an anti-Xyn1 antiserum and by xylanase assays (52), respectively.

Amino acid sequencing, xylanase assays, and determination of protein concentrations. The amino-terminal sequence was determined using an ABI model 491 protein sequencer (Applied Biosystems, Foster City, CA) as described previously (52). Xyn1 activity was measured using water-soluble xylan as a substrate, and 1 U of the enzyme activity was defined as the amount of enzyme that liberated 1 µmol of xylose per min from the substrate (52). Protein concentrations were determined by the method of Bradford (5) using bovine serum albumin as the standard.

Nucleotide sequence accession numbers. The 12-kbp nucleotide sequence of the xyn1 locus has been deposited in the DDBJ/EMBL/GenBank databases under accession no. AB274730.


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RESULTS
 
LppX is a membrane lipoprotein in Paenibacillus sp. strain W-61. We determined a 12-kbp flanking DNA region of xyn1 by inverse PCR walking based on the known xyn1 sequence (52). This DNA region contained four (orf1 through orf4) and three (lppX, orf5, and orf6) putative genes upstream and downstream of xyn1, respectively (Fig. 1). Orfs 1, 2, 3, 4, 5, and 6 exhibited 26, 77, 29, 47, 89, and 68% identity with the endo-β-1,4-glucanase of Clostridium cellulovorans (accession no. AAB40891), an intracellular exo-oligoxylanase of Bacillus halodurans C-125 (16, 22), Cryptococcus neoformans var. grubii CAS35p protein, which is involved in the formation of the glucuronoxylomannan capsule (7, 37), the feruloyl esterase domain of Clostridium thermocellum endo-β-1,4-xylanase (3), the endo-1,3-1,4-β-glucanases of Paenibacillus polymyxa (AAN85721) and Bacillus macerans (4), and a substrate-binding protein of an ABC sugar transporter of Bacillus clausii (BAD63263), respectively. The products of these open reading frames were presumed to be xylan utilization-related proteins. In contrast, lppX, immediately downstream of xyn1, encoded a 279-amino-acid protein that shared 61% amino acid identity with the XaiF of Geobacillus stearothermophilus (8, 9). We found that the LppX signal peptide sequence contained a tetrapeptide identical to a lipobox (Leu-Ser-Ala-Cys) that is conserved among bacterial membrane lipoproteins (53) (see Fig. S1 in the supplemental material). A pre-form of lipoprotein is cleaved by signal peptidase II between the Ala and Cys residues in the lipobox, and the amino-terminal Cys residue of a mature lipoprotein is subsequently modified with diacylglycerol and a fatty acid via thioester and amide linkages, respectively (53). Because the lines of evidence presented below support the notion that this gene encodes a lipoprotein that is essential for Xyn1 production, we refer to it as lppX (for "lipoprotein gene required for Xyn1 production").


Figure 1
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  		FIG. 1. Gene map of the xyn1 locus of Paenibacillus sp. strain W-61. Open, filled, and hatched arrows indicate the directions and sizes of the potential genes orf1 through orf6, xyn1, and lppX, respectively. The bent arrow in front of orf3 represents a putative promoter of the orf3-lppX operon.

To prove that LppX is a membrane lipoprotein, we initially tried to detect [14C]palmitic acid-labeled LppX in wild-type cells but were unable to do so, presumably because the amounts of this lipoprotein in cells were very low. We therefore cloned the Pxyn1::lppX fusion gene into the high-copy-number plasmid vector pHY300PLK (ca. 200 copies per chromosome) and introduced the resultant plasmid, pHPX4T (lppX), into strain PSC301 (lppX::cat). We then grew strains PSC301/pHY300PLK (control) and PSC301/pHPX4T (lppX) and analyzed LppX and 14C-labeled proteins in their cytosols and cytoplasmic membranes. Western blotting using an anti-LppX antiserum identified the 31-kDa LppX protein in the cytoplasmic membranes of the PSC301/pHPX4T cells (Fig. 2A). No LppX protein was detected in the cytosolic proteins of the PSC301/pHPX4T cells or in either the cytosolic proteins or the cytoplasmic membranes of the lppX-negative PSC301 cells (Fig. 2A). [14C]palmitic acid was incorporated into a 31-kDa protein and into five other, larger proteins in the PSC301/pHPX4T membranes (Fig. 2B). The five radioactive proteins of >31 kDa were also present in the cytoplasmic membranes of the lppX-negative cells (Fig. 2B). These findings support the notion that LppX is a fatty acylated protein that is located predominantly in the cytoplasmic membranes.


Figure 2
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  		FIG. 2. Labeling of LppX with [14C]palmitic acid in vivo. PSC301 (lppX::cat) cells carrying either the pHY300PLK vector or plasmid pHPX4T (pHY300PLK::lppX) were grown in Medium I containing [14C]palmitic acid to the late-exponential-growth phase. The envelopes and cytosolic proteins (1 mg each) of the cells were resolved by SDS-PAGE in order to detect LppX by Western blotting (A) and 14C-labeled proteins by using a FIA-2000 fluoroimaging analyzer (B). Arrowheads indicate the 31-kDa LppX protein (A) and the position corresponding to 31 kDa (B). Lanes 1, PSC301 soluble proteins; lanes 2, PSC301 cell envelope proteins; lanes 3, PSC301/pHPX4T soluble proteins; lanes 4, PSC301/pHPX4T cell envelope proteins.

LppX is located on the outer surface of the cytoplasmic membrane in Paenibacillus sp. strain W-61. To determine whether LppX is located in the inner or outer layers of the cytoplasmic membrane, we prepared protoplasts and inside-out vesicles of the cytoplasmic membranes of PSC301/pHPX4T. Proteinase K rapidly and completely digested LppX molecules in the protoplasts (Fig. 3A), whereas those in the inside-out membrane vesicles escaped hydrolysis. However, they were digested by proteinase K when the membrane barriers were collapsed by the addition of 1% (wt/vol) SDS (Fig. 3B). Thus, LppX appeared to be located on the outer layer of the cytoplasmic membranes.

Xyn1 synthesis requires lppX in Paenibacillus sp. strain W-61. To examine whether lppX is indispensable for Xyn1 synthesis, we cultured strains W-61 (wild type), PSC301 (lppX::cat)/pHY300PLK, and PSC301/pHPX4T in Medium I supplemented with water-soluble xylan as a carbon source (inducible conditions). The growth of PSC301/pHY300PLK was remarkably slower than that of the other strains. Quantitative analysis of Xyn1 in the supernatants of late-exponential-growth-phase cells (1 x 109/ml) by Western blotting against a standard curve constructed with purified Xyn1 revealed that PSC301/pHY300PLK formed only 0.4 ng Xyn1 per ml of culture, whereas the wild-type strain produced 8 ng of the enzyme per ml of culture. Plasmid pHPX4T carrying lppX restored Xyn1 synthesis by the lppX mutant to half of the wild-type level (4 ng/ml culture). Both PSC301/pHY300PLK and PSC301/pHPX4T generated Xyn3 and Xyn5 in amounts comparable to those produced by the wild-type strain (data not shown). Thus, lppX appeared to be importantly and specifically required for Xyn1 synthesis by Paenibacillus sp. strain W-61.

LppX is required to produce active Xyn1 in E. coli. To further examine the participation of lppX in Xyn1 production, we introduced plasmid pXFT, containing both xyn1 and lppX, or plasmid pX1T, harboring only xyn1, into heterologous E. coli DH5{alpha} cells. E. coli DH5{alpha}/pXFT formed large and clear halos around its colonies on 0.5% (wt/vol) Remazol Brilliant Blue R-D-xylan plates, whereas E. coli DH5{alpha}/pX1T formed small and cloudy halos (data not shown). However, as quantified by Western blotting E. coli DH5{alpha}/pX1T synthesized about half the amount of Xyn1 protein produced by E. coli DH5{alpha}/pXFT (23 versus 50 ng/109 cells), implying that DH5{alpha}/pX1T synthesizes Xyn1, but in an inactive form. Thus, we measured the enzyme activities of Xyn1 from E. coli DH5{alpha}/pX1T (xyn1) and Xyn1 from E. coli DH5{alpha}/pXFT (xyn1 lppX). E. coli DH5{alpha}/pXFT produced 1.8 U of Xyn1 in culture (100 ml), and most activity (95%) was detected in the soluble-protein fraction (Fig. 4). E. coli DH5{alpha}/pX1T cells produced a quantity of Xyn1 equivalent to 65% of the amount generated by strain DH5{alpha}/pXFT (Fig. 4). However, Xyn1 activity was undetectable in DH5{alpha}/pX1T cell extracts, and almost all the Xyn1 protein was located in the cell envelope fraction (Fig. 4), indicating that LppX is required for the production of soluble and active Xyn1 but not for xyn1 transcription or translation.


Figure 4
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  		FIG. 4. lppX is indispensable for the synthesis of active Xyn1 in E. coli. Extracts (C) of E. coli DH5{alpha} cells harboring plasmid pX1T (xyn1) or pXFT (xyn1-lppX) growing exponentially in LB medium were separated into soluble proteins (S) and cell envelopes (E) by centrifugation at 100,000 x g for 120 min. The amounts of Xyn1 in these cell fractions were determined by Western blotting using a standard curve constructed with purified Xyn1 (52): the signal intensity of Xyn1 was proportional at least up to 100 ng. The amounts of Xyn1 in each sample are presented as ratios (percentages) of the amount in the DH5{alpha}/pXFT cell extract. Xylanase activities were measured using water-soluble xylan as a substrate and are expressed in units.

Location of LppX in E. coli membranes. To ascertain the membrane location of LppX expressed in E. coli cells, we determined by Western blotting the amounts of LppX in soluble proteins of the cytosol, the periplasm, the inner membranes, and the outer membranes of E. coli DH5{alpha}/pXFT (xyn1 lppX) cells growing exponentially in LB broth (see Materials and Methods) (Fig. 5A). LppX was predominantly (~90%) located in the outer membranes, with very small quantities (~5% each) in the soluble proteins and inner membranes (Fig. 5A). To determine the location of LppX on the outer membrane, we digested the intact cells and inside-out vesicles of the outer membranes of E. coli DH5{alpha}/pXFT with proteinase K. The proteinase reduced the levels of several proteins in the E. coli cells (including outer membrane protein OmpA) that are exposed on the cell surface (27), whereas it only marginally affected the amounts of LppX (Fig. 5B). In the presence of 1% SDS, the proteinase completely digested LppX in the cells (data not shown). In addition, the proteinase completely digested LppX in the inside-out vesicles of the outer membranes (Fig. 5B), confirming that LppX is located on the inner layer of the outer membranes, with its entire polypeptide chain exposed to the periplasm.

Destinations of LppX mutants in E. coli membranes. The amino acid residues at positions +2 (Asp) and +3 (Asp, Arg, or Gln) relative to the lipobox Cys residue (position +1) of E. coli lipoproteins serve as signals for sorting to the inner or the outer membranes (24, 34, 54). To examine whether the membrane destination of LppX can be manipulated by modifying sorting-signal amino acids, we constructed three mutant lppX genes—lppX(C19A), lppX(S20D), and lppX(S20D S21D)—and cloned them into plasmid pUC119 to create plasmids pC19A, pS20D, and pS20DS21D, respectively (see Materials and Methods). We examined the cellular locations of LppX in E. coli DH5{alpha} harboring a mutant lppX gene on a plasmid by Western blotting. LppX(C19A) was detected predominantly in the periplasm (Fig. 5C). LppX(S20D) was located in the outer membranes, like wild-type LppX (data not shown). This signal alone appeared to be insufficient to retain LppX in the inner membranes. However, LppX(S20D S21D), which had a canonical Asp-Asp signal for sorting to the inner membrane, was located mostly in the inner membranes (Fig. 5D).

Activity and cellular location of Xyn1 synthesized in the presence or absence of lppX in E. coli. In the presence of LppX, Xyn1 is secreted into the medium in an active form, whereas it sediments together with cell envelopes after centrifugation without LppX (Fig. 4). However, whether the inactive Xyn1 was insoluble or was associated with the cell membranes remained obscure, so we expressed Xyn1 in E. coli DH5{alpha}/pX1T (xyn1) and DH5{alpha}/pXFT (xyn1 lppX) and then prepared the spheroplasts and periplasmic proteins. Xyn1 in E. coli DH5{alpha}/pX1T (xyn1) was detected in the spheroplasts, whereas in E. coli DH5{alpha}/pXFT (xyn1 lppX), Xyn1 was detected mostly in the periplasm (Fig. 6A). Xyn1 in the E. coli DH5{alpha}/pXFT periplasm was active (180 U/mg protein). However, the xylanase in the DH5{alpha}/pX1T spheroplasts exhibited no measurable activity (<1 U/mg protein), although the amount was equivalent to 12.6% of the amount of xylanase in the DH5{alpha}/pXFT periplasm (Fig. 6A). Furthermore, we separated the inner and outer membranes from E. coli DH5{alpha}/pX1T cells by sucrose density gradient centrifugation (35) and examined the presence of Xyn1 by Western blotting. Xyn1 was located only in the inner membrane and was found nowhere in the outer membrane or cytosol (data not shown). The amino-terminal sequence of the inactive Xyn1 in DH5{alpha}/pX1T cells was ATSSAA, which was consistent with that for the native rXyn1 (52), indicating that inactive Xyn1 had been processed by a signal peptidase between residues 28 and 29. Based on these results, we postulated that Xyn1 remains in the inner membranes as an inactive form without LppX.

When the DH5{alpha}/pX1T spheroplasts were incubated with proteinase K, the 21-kDa Xyn1 protein was digested into a 19-kDa polypeptide (Fig. 6B), suggesting that a digested 2-kDa region is exposed outside the spheroplasts and that another region of 19 kDa resides inside the spheroplasts or within the inner membrane. To determine the location of the 19-kDa polypeptide in the DH5{alpha}/pX1T spheroplasts, we prepared inside-out membrane vesicles of the spheroplasts and digested them with proteinase K. The proteinase segmented Xyn1 into small fragments (<0.8 kDa) (Fig. 6B) Thus, in the absence of LppX, Xyn1 seems unable to complete its secretion processes and pauses in the inner membranes. This, in turn, exposes the 2-kDa amino-terminal polypeptide to the periplasm and the remaining 19-kDa carboxyl-terminal polypeptide to the cytoplasm.

LppX liberates inactive secretion intermediates of Xyn1 from the inner membranes as active enzymes. LppX(C19A) resided freely in the periplasm (Fig. 5C), and LppX(S20D S21D) was anchored to the inner membranes (Fig. 5D). To examine whether or not E. coli DH5{alpha}/pC19A and E. coli DH5{alpha}/pS20DS21D produce active Xyn1 as well as wild-type LppX from E. coli DH5{alpha}/pXFT (Fig. 4), we compared the amounts of Xyn1 these strains produced with that produced by E. coli DH5{alpha}/pXFT. Both E. coli DH5{alpha}/pC19A and E. coli DH5{alpha}/pS20DS21D produced active Xyn1 in the periplasm at levels (134 and 167 U/mg protein, respectively) comparable to that produced by E. coli DH5{alpha}/pXFT (1.8 U/100 ml of culture) (Fig. 7A). Western blotting showed that E. coli DH5{alpha}/pC19A and E. coli DH5{alpha}/pS20DS21D produced Xyn1 exclusively in the periplasm (Fig. 7A), indicating that both the free and anchored forms of LppX are functional in terms of active Xyn1 production.


Figure 7
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  		FIG. 7. (A) Xyn1 secretion in E. coli DH5{alpha}/pC19A and DH5{alpha}/pS20DS21D. (A) Spheroplasts (S) and periplasms (P) were prepared from E. coli DH5{alpha}/pC19A and DH5{alpha}/pS20DS21D. The amounts of Xyn1 in the spheroplasts (secretion intermediates) and periplasms (secreted Xyn1) from 5 x 106 cells of each strain were quantified by Western blotting. (B) Chaperone activity of LppX from E. coli BL21(DE3)/pC19A2. E. coli DH5{alpha}/pX1T spheroplasts (5 x 108) containing inactive Xyn1 were incubated without (left) or with (right) the periplasmic proteins (200 µg) from isopropyl-β-D-thiogalactopyranoside (IPTG)-induced E. coli BL21(DE3)/pC19A2 in 0.85-ml reaction mixtures (see Materials and Methods) at 30°C for 10 min. After incubation, spheroplasts were collected by centrifugation at 4°C and 10,000 x g for 30 min in order to determine the amount of Xyn1 that remained bound to the spheroplasts (S) and the amount that was free in the supernatant buffer (F) by Western blotting.

Function of LppX as a molecular chaperone in Xyn1 secretion. Xyn1 cannot cross the inner membranes in the absence of LppX and remains within the membranes without proceeding to the periplasm (Fig. 6A). Since free LppX(C19A) can help secretion intermediates of Xyn1 to complete passage through the inner membranes in E. coli DH5{alpha}(pC19A) (Fig. 7A), we examined whether free LppX(C19A) can rescue the secretion intermediates in the inner membranes of DH5{alpha}/pX1T spheroplasts as active enzymes in vitro. When DH5{alpha}/pX1T spheroplasts were incubated with BL21(DE3)/pC19A2 periplasmic proteins, which contained LppX(C19A), a significant amount of Xyn1 was liberated from the spheroplasts into the solution (Fig. 7B, right) as an active enzyme (450 U/mg protein).In contrast, when the DH5{alpha}/pX1T spheroplasts were incubated with periplasmic proteins of the control strain BL21(DE3)/pET-15b without LppX(C19A), most (90%) of the Xyn1 remained in the spheroplasts (Fig. 7B, left). The small amount of Xyn1 released into the reaction mixture during incubation had no enzyme activity (<1 U/mg protein). These results indicate that LppX liberates secretion intermediates of Xyn1 from the inner membranes as active enzymes. Hence, it was demonstrated that LppX functions as a molecular chaperone in Xyn1 secretion.


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DISCUSSION
 
LppX is structurally similar (61% identity) to Geobacillus stearothermophilus XaiF, which positively regulates the expression of xynA (encoding a xylanase A of GH family 11 with 85% identity to Xyn1 [8, 9]) by protecting xynA transcripts from digestion by 3'-to-5' RNase through binding to their 3' untranslated regions (26). LppX apparently is not involved in xyn1 mRNA stability, because the amounts of xyn1 mRNA were comparable between lppX mutants with and without pHPX4T, carrying lppX (see Fig. S3 in the supplemental material), and between E. coli DH5{alpha}/pX1T (xyn1) and E. coli DH5{alpha}/pXFT (xyn1-lppX) (data not shown). LppX and XaiF each have a tetrapeptide identical to a lipobox (Leu-Ser-Ala-Cys) that is conserved among bacterial membrane lipoproteins (53) (see Fig. S2 in the supplemental material). However, LppX and XaiF differ at their amino termini. LppX has two positively charged amino acids after the amino-terminal methionine (M-K-K) for the interaction with the cytoplasmic membranes during an initial stage of secretion processes, as well as a following hydrophobic peptide (M-L-F-L-F-I-A-A-V-A), a structural feature of signal peptides. In contrast, XaiF has no positively charged amino acid residue at the relevant positions (M-S-M-I) (see Fig. S2 in the supplemental material). Thus, XaiF may be a cytoplasmic protein, in accordance with its role in the protection of the xynA transcripts from RNase (26). The cellular location of XaiF remains to be determined.

LppX can be labeled in vivo with exogenous [14C]palmitic acid (Fig. 2B), and it is located on the outer surface of the cytoplasmic membrane (Fig. 3B), in agreement with the notion that LppX is a lipoprotein. An LppX mutant in which the 19th cysteine, a plausible lipomodification residue in the lipobox (see Fig. S2 in the supplemental material), was replaced with alanine was exported into the periplasm but was not anchored to the cytoplasmic membrane. Nascent LppX, therefore, is likely cleaved by a signal peptidase in front of the 19th cysteine residue (see Fig. S1 in the supplemental material); then the cysteine would be modified with fatty acids and diacylglycerol to anchor mature LppX to the outer leaflet of the cytoplasmic membrane at the amino terminus. In E. coli, wild-type LppX is delivered to the inner layer of the outer membrane (Fig. 5A and B), probably because the second and third amino-terminal residues are serines, which serve as a sorting signal for the outer membrane in gram-negative bacteria (54). In fact, LppX(S20D S21D), with conversions of the 20th and 21st serines to aspartate residues, the canonical signal for sorting to the inner membrane (54), was found predominantly in the inner membranes (Fig. 5D). These results demonstrate that LppX has the sorting property of a lipoprotein (50, 54), and they substantiate the notion that LppX is a lipoprotein.

According to Ellis (14), molecular chaperones are defined as a functional class of unrelated families of proteins that assist in the correct noncovalent assembly of other structures containing polypeptides in vivo but are not components of the assembled structures when performing their normal biological functions. The correct assembly of several cytosolic proteins under normal conditions and the refolding of unfolded cytosolic proteins induced by exposure to stresses such as high temperatures require molecular chaperones (18, 33). Two major classes of molecular chaperones, GroEL-GroES and DnaJ-DnaK complexes, assist the folding and refolding of cytosolic proteins (33). They also help to maintain the unfolded state of secreted proteins before and during secretion processes (33, 51). SecB and the signal recognition particle SRP serve as molecular chaperones for proteins secreted via the Sec-dependent pathway. They bind to nascent polypeptides and guide them to the secretion apparatus (33). Some effector proteins that are secreted via the type III secretion system require the help of cognate effector chaperones for translocation from the bacterial cytosol into the host cell interior (15, 19). Other examples of molecular chaperones are the Lif and LimA proteins of Burkholderia glumae and Burkholderia cepacia, respectively (13, 21, 42). These are inner membrane proteins with a large C-terminal region protruding into the periplasm, and they specifically help cognate lipases to assume the active conformation after export through the inner membrane (13, 21, 42).

Besides molecular chaperones, the membrane lipoprotein peptidyl prolyl-cis/trans isomerase (PPI; EC 5.2.1.89) builds up active folds of secreted proteins by converting cis/trans configurations of proline residues and hence is essential for some proteins to assume the active form after secretion (28); a prsA mutant of B. subtilis lacking PPI is defective in the secretion of {alpha}-amylase and other proteins (29, 30). LppX is structurally different from general secretion proteins such as SecYEG (36) and chaperones of proteins secreted via the type III secretion system (15, 19).

In the absence of LppX, Xyn1 is normally transcribed, translocated into the periplasm, and processed into a mature form but not secreted. The presence of significant amounts of Xyn1 in DH5{alpha}/pXFT (xyn1-lppX) spheroplasts (~10% in the periplasm) (Fig. 6A) implies that the process of secretion across the cytoplasmic membranes is a rate-limiting step in Xyn1 production and that Xyn1 requires the function of LppX to complete the secretion process. Since LppX can act on Xyn1 secretion intermediates from outside the membranes and the secretion intermediates expose the 2-kDa amino-terminal polypeptide of Xyn1 (Fig. 6B), the lipoprotein might interact with this amino-terminal polypeptide so that the secretion intermediates form the fold necessary for detachment from the cytoplasmic membranes.

The signal peptide of Xyn1 can be removed in the absence of LppX, indicating that the lipoprotein is not involved in the processing of Xyn1 during secretion. Clearly, LppX is not a component of active Xyn1, and it appears not to modify the covalent assembly of Xyn1. Finally, LppX can activate and liberate inactive Xyn1 on the protoplast membranes (Fig. 7B), thus fulfilling the definition of a molecular chaperone (14). LppX has no significant similarity either to PPI or to any known bacterial molecular chaperones. An lppX mutant produced Xyn3 and Xyn5 normally, and lppX was specifically expressed together with xyn1 in the presence of xylan (see Fig. S3 in the supplemental material). To our knowledge, LppX is a novel molecular chaperone lipoprotein specific to Xyn1. LppX has the DUF232 domain (Pfam 03781) between residues 50 and 276. This domain is found in diverse bacterial and eukaryotic proteins, including formylglycine-generating enzyme (FGE), which catalyzes the posttranscriptional formation of the C{alpha}-formylglycine residue at the active site of eukaryotic sulfate esterase (43, 47). The paralogue of FGE, pFGE, which also has the DUF232 domain but not FGE activities (11), is required by FGE for this activity. pFGE is a dimer with a deep cleft for binding a sulfatase peptide bearing a recognition motif for FGE. Presumably, pFGE forms a heterodimer with FGE bound to sulfatase during the formylation of glycylglycine at the active site (11). LppX might bind to the amino-terminal 2-kDa polypeptides of Xyn1 secretion intermediates via the DUF232 domain to induce a conformational change in the intermediates that allows their escape from the cytoplasmic membrane.

While xylanases of GH family 11 are highly conserved and are distributed among gram-positive and gram-negative bacteria (10), LppX paralogues (score, >300) occur in some gram-positive bacteria, such as Bacillus halodurans (49), Clostridium phytofermentans (accession no. ABX4272), Clostridium beijerinckii (ABR35179), and Geobacillus stearothermophilus (8). They all have a tetrapeptide [(L/V/A)-(S/T/G)-A-C] analogous to the lipobox (L-S-A-C) of LppX at the relevant amino-terminal region (see Fig. S2 in the supplemental material), and like lppX, they are located downstream of a GH family 11 xylanase gene. They might play a role similar to that of LppX in the production of cognate family 11 xylanases. Bacillus subtilis 168 actively produces a XynA of xylanase family 11 that has 79% identity (87% similarity) with Xyn1, but this strain has no lppX counterpart (32, 44, 48). Xyn1 might lack a polypeptide region necessary for correct self-folding and active conformation, or B. subtilis 168 might have a protein that is structurally dissimilar but functionally similar to LppX. These issues remain to be elucidated.


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ACKNOWLEDGMENTS
 
This work was supported in part by the Pioneering Research Project in Biotechnology of the Ministry of Agriculture, Forestry, and Fisheries of Japan (to Y.K.).


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Human Health and Nutrition, Graduate School of Comprehensive Human Sciences, Shokei Gakuin University, Yurigaoka 4-10-1, Natori 981-1295, Japan. Phone and fax: 81-22-381-3347. E-mail: ykamio{at}shokei.ac.jp Back

{triangledown} Published ahead of print on 22 December 2008. Back

{dagger} Supplemental material for this article may be found at http://jb.asm.org/. Back

{ddagger} M.F. and S.W. contributed equally to this work. Y. Itoh passed away on 4 October 2008. Back

§ Present address: Central Research Institute, Mizkan Group Co. Ltd., Handa 475-8585, Japan. Back


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REFERENCES
 
    1
  1. Aiba, H., S. Adhya, and B. de Crombrugghe. 1981. Evidence for two functional gal promoters in intact Escherichia coli cells. J. Biol. Chem. 256:11905-11910.[Abstract/Free Full Text]
    2. 2
  2. Beg, Q. K., M. Kapoor, L. Mahajan, and G. S. Hoondal. 2001. Microbial xylanases and their applications: a review. Appl. Microbiol. Biotechnol. 56:326-338.[CrossRef][Medline]
    3. 3
  3. Blum, D. L., I. R. Kataeva, X.-L. Li, and L. G. Ljungdahl. 2000. Feruloyl esterase activity of the Clostridium thermocellum cellulosome can be attributed to previously unknown domains of XynY and XynZ. J. Bacteriol. 182:1346-1351.[Abstract/Free Full Text]
    4. 4
  4. Borriss, R., K. Buettner, and P. Maentsaelae. 1990. Structure of the β-1,3-1,4-glucanase gene of Bacillus macerans: homologies to other β-glucanases. Mol. Gen. Genet. 222:278-283.[CrossRef][Medline]
    5. 5
  5. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254.[CrossRef][Medline]
    6. 6
  6. Calvin, N. M., and P. C. Hanawalt. 1988. High-efficiency transformation of bacterial cells by electroporation. J. Bacteriol. 170:2796-2801.[Abstract/Free Full Text]
    7. 7
  7. Chang, Y. C., L. A. Penoyer, and K. J. Kwon-Chung. 1996. The second capsule gene of Cryptococcus neoformans, CAP64, is essential for virulence. Infect. Immun. 64:1977-1983.[Abstract]
    8. 8
  8. Cho, S. G., and Y. J. Choi. 1995. Nucleotide sequence analysis of an endo-xylanase gene (xynA) from Bacillus stearothermophilus. J. Microbiol. Biotechnol. 5:117-124.[CrossRef]
    9. 9
  9. Cho, S. G., and Y. J. Choi. 1998. Characterization of the xaiF gene encoding a novel xylanase-activity-increasing factor, XaiF. J. Microbiol. Biotechnol. 8:378-387.
    10. 10
  10. Collins, T., C. Gerday, and G. Feller. 2005. Xylanases, xylanase families and extremophilic xylanases. FEMS Microbiol. Rev. 29:2-23.
    11. 11
  11. Dickmanns, A., B. Schmidt, M. G. Rudolph, M. Mariappan, T. Dierks, K. van Figura, and R. Ficner. 2005. Crystal structure of human pFGE, the paralog of the C{alpha}-formylglycine-generating enzyme. J. Biol. Chem. 250:15180-15187.
    12. 12
  12. Egelseer, E., I. Schocher, M. Sara, and U. B. Sleytr. 1995. The S-layer from Bacillus stearothermophilus DSM 2358 functions as an adhesion site for a high-molecular-weight amylase. J. Bacteriol. 177:1444-1451.[Abstract/Free Full Text]
    13. 13
  13. El Khattabi, M., P. Van Gelder, W. Bitter, and J. Tommassen. 2000. Role of the lipase-specific foldase of Burkholderia glumae as a steric chaperone. J. Biol. Chem. 275:26885-26891.[Abstract/Free Full Text]
    14. 14
  14. Ellis, R. J. 1993. The general concept of molecular chaperones. Philos. Trans. R. Soc. Lond. B 339:257-261.[Abstract/Free Full Text]
    15. 15
  15. Feldman, M. F., and G. R. Cornelis. 2003. The multitalented type III chaperones: all you can do with 15 kDa. FEMS Microbiol. Lett. 219:151-158.[CrossRef][Medline]
    16. 16
  16. Fushinobu, S., M. Hidaka, Y. Honda, T. Wakagi, H. Shoun, and M. Kitaoka. 2005. Structural basis for the specificity of the reducing end xylose-releasing exo-oligoxylanase from Bacillus halodurans C-125. J. Biol. Chem. 280:17180-17186.[Abstract/Free Full Text]
    17. 17
  17. Futai, M. 1974. Orientation of membrane vesicles from Escherichia coli prepared by different procedures. J. Membrane Biol. 15:15-28.[CrossRef][Medline]
    18. 18
  18. Genevaux, P., C. Georgopoulos, and W. L. Kelley. 2007. The Hsp70 chaperone machines of Escherichia coli: a paradigm for the repartition of chaperone functions. Mol. Microbiol. 66:840-857.[CrossRef][Medline]
    19. 19
  19. Ghosh, P. 2004. Process of protein transport by the type III secretion system. Microbiol. Mol. Biol. Rev. 68:771-795.[Abstract/Free Full Text]
    20. 20
  20. Higuchi, R., B. Krummel, and R. K. Sasaki. 1988. A general method of in vitro preparation and specific mutagenesis of DNA fragments: study of protein and DNA interactions. Nucleic Acids Res. 16:7351-7367.[Abstract/Free Full Text]
    21. 21
  21. Hobson, A. H., C. M. Buckley, J. L. Aamand, S. T. Jorgensen, B. Diderichsen, and D. J. McConnell. 1993. Activation of a bacterial lipase by its chaperone. Proc. Natl. Acad. Sci. USA 90:5682-5686.[Abstract/Free Full Text]
    22. 22
  22. Honda, Y., and M. Kitamoto. 2004. A family 8 glycoside hydrolase from Bacillus halodurans C-125 (BH2105) is a reducing end xylose-releasing exo-oligoxylanase. J. Biol. Chem. 279:55097-55103.[Abstract/Free Full Text]
    23. 23
  23. Horinouchi, S., and B. Weisblum. 1982. Nucleotide sequence and functional map of pC194, a plasmid that specifies inducible chloramphenicol resistance. J. Bacteriol. 150:815-825.[Abstract/Free Full Text]
    24. 24
  24. Inouye, S., G. Duffaud, and M. Inouye. 1986. Structural requirement at the cleavage site for efficient processing of the lipoprotein secretory precursor of Escherichia coli. J. Biol. Chem. 261:10970-10975.[Abstract/Free Full Text]
    25. 25
  25. Ito, Y., T. Tomota, N. Roy, A. Nakano, N. Sugawara-Tomita, S. Watanabe, N. Okai, N. Abe, and Y. Kamio. 2003. Cloning, expression, and cell surface localization of Paenibacillus sp. strain W-61 xylanase 5, a multidomain xylanase. Appl. Environ. Microbiol. 69:6969-6978.[Abstract/Free Full Text]
    26. 26
  26. Jeong, M. Y., E. R. Lee, C. W. Yun, S. G. Cho, and Y. J. Choi. 2006. Post-transcriptional regulation of the xynA expression by a novel mRNA binding protein, XaiF. Biochem. Biophys. Res. Commun. 351:153-158.[CrossRef][Medline]
    27. 27
  27. Kamio, Y., and H. Nikaido. 1976. Outer membrane of Salmonella typhimurium: accessibility of phospholipid head groups to phospholipase C and cyanogen bromide activated dextran in the external medium. Biochemistry 15:2561-2570.[CrossRef][Medline]
    28. 28
  28. Kim, J. H., I. S. Park, and B. G. Kim. 2005. Development and characterization of membrane surface display system using molecular chaperon, PrsA, of Bacillus subtilis. Biochem. Biophys. Res. Commun. 334:1248-1253.[CrossRef][Medline]
    29. 29
  29. Kontinen, V. P., and M. Sarvas. 1988. Mutants of Bacillus subtilis defective in protein export. J. Gen. Microbiol. 134:2333-2344.[Abstract/Free Full Text]
    30. 30
  30. Kontinen, V. P., and M. Sarvas. 1993. The PrsA lipoprotein is essential for protein secretion in Bacillus subtilis and sets a limit for high-level secretion. Mol. Microbiol. 8:727-737.[Medline]
    31. 31
  31. Kulkarni, N., A. Shendye, and M. Rao. 1999. Molecular and biotechnological aspects of xylanases. FEMS Microbiol. Rev. 23:411-456.[CrossRef][Medline]
    32. 32
  32. Kunst, F., N. Ogasawara, I. Moszer, A. M. Albertini, G. Alloni, et al. 1997. The complete genome sequence of the gram-positive bacterium Bacillus subtilis. Nature 390:249-256.[CrossRef][Medline]
    33. 33
  33. Lund, P. A. 2001. Microbial molecular chaperones. Adv. Microb. Physiol. 44:93-140.[Medline]
    34. 34
  34. Matsuyama, S., T. Tajima, and H. Tokuda. 1995. A novel periplasmic carrier protein involved in the sorting and transport of Escherichia coli lipoproteins destined for the outer membrane. EMBO J. 14:3365-3372.[Medline]
    35. 35
  35. Miura, T., and S. Mizushima. 1968. Separation by density gradient centrifugation of two types of membranes from spheroplast membrane of Escherichia coli K-12. Biochim. Biophys. Acta 150:159-161.[Medline]
    36. 36
  36. Mori, H., and K. Ito. 2001. The Sec protein-translocation pathway. Trends Microbiol. 9:494-500.[CrossRef][Medline]
    37. 37
  37. Moyrand, F., T. Fontaine, and G. Janbon. 2007. Systematic capsule gene disruption reveals the central role of galactose metabolism on Cryptococcus neoformans virulence. Mol. Microbiol. 64:771-781.[CrossRef][Medline]
    38. 38
  38. Nguyen, V. D., Y. Kamio, N. Abe, J. Kaneko, and K. Izaki. 1991. Purification and properties of β-1,4-xylanase from Aeromonas caviae W-61. Appl. Environ. Microbiol. 57:445-449.[Abstract/Free Full Text]
    39. 39
  39. Nguyen, V. D., Y. Kamio, N. Abe, J. Kaneko, and K. Izaki. 1993. Purification and properties of β-1,4-xylanases 2 and 3 from Aeromonas caviae W-61. Biosci. Biotechnol. Biochem. 57:1708-1712.
    40. 40
  40. Okai, N., M. Fukasaku, J. Kaneko, T. Tomita, K. Muramoto, and Y. Kamio. 1998. Molecular properties and activity of carboxyl-terminal truncated form of xylanase 3 from Aeromonas caviae W-61. Biosci. Biotechnol. Biochem. 62:1560-1567.[CrossRef][Medline]
    41. 41
  41. Osborn, M. J., J. E. Gander, E. Parisi, and J. Carson. 1972. Mechanism of assembly of the outer membrane of Salmonella typhimurium. Isolation and characterization of cytoplasmic and outer membrane. J. Biol. Chem. 247:3962-3972.[Abstract/Free Full Text]
    42. 42
  42. Pauwels, K., A. Lustig, L. Wyns, J. Tommassen, S. N. Savvides, and P. Van Gelder. 2006. Structure of a membrane-based steric chaperone in complex with its lipase substrate. Nat. Struct. Mol. Biol. 13:374-375.[CrossRef][Medline]
    43. 43
  43. Recksiek, M., T. Selmer, T. Dierks, B. Schmidt, and K. von Figura. 1998. Sulfatases, trapping of the sulfated enzyme intermediate by substituting the active site formylglycine. J. Biol. Chem. 273:6096-6103.[Abstract/Free Full Text]
    44. 44
  44. Roncero, M. I. 1983. Genes controlling xylan utilization by Bacillus subtilis. J. Bacteriol. 156:257-263.[Abstract/Free Full Text]
    45. 45
  45. Roy, N., N. Okai, T. Tomita, K. Muramoto, and Y. Kamio. 2000. Purification and some properties of high-molecular-weight xylanases, the xylanase 4 and 5 of Aeromonas caviae W-61. Biosci. Biotechnol. Biochem. 64:408-413.[CrossRef][Medline]
    46. 46
  46. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
    47. 47
  47. Schmidt, B., T. Selmer, A. Ingendoh, and K. von Figura. 1995. A novel amino acid modification in sulfatases that is defective in multiple sulfatase deficiency. Cell 82:271-278.
    48. 48
  48. St John, F. J., J. D. Rice, and J. F. Preston. 2006. Characterization of XynC from Bacillus subtilis subsp. subtilis strain 168 and analysis of its role in depolymerization of glucuronoxylan. J. Bacteriol. 188:8617-8626.[Abstract/Free Full Text]
    49. 49
  49. Takami, H., K. Nakasone, Y. Takaki, G. Maeno, R. Sasaki, N. Masui, F. Fuji, C. Hirama, Y. Nakamura, S. Kuhara, and K. Horikoshi. 2000. Complete genome sequence of the alkaliphilic bacterium Bacillus halodurans and genomic sequence comparison with Bacillus subtilis. Nucleic Acids Res. 28:4317-4331.[Abstract/Free Full Text]
    50. 50
  50. Terada, M., T. Kuroda, S. Matsuyama, and H. Tokuda. 2001. Lipoprotein sorting signals evaluated as the LolA-dependent release of lipoproteins from the cytoplasmic membrane of Escherichia coli. J. Biol. Chem. 276:47690-47694.[Abstract/Free Full Text]
    51. 51
  51. van Wely, K. H. M., J. Swaving, R. Freudl, and A. J. M. Driessen. 2001. Translocation of proteins across the cell envelope of Gram-positive bacteria. FEMS Microbiol. Rev. 25:437-454.[Medline]
    52. 52
  52. Watanabe, S., V. D. Nguyen, J. Kaneko, Y. Kamio, and S. Yoshida. 2008. Cloning, expression, and transglycosylation reaction of Paenibacillus sp. strain W-61 xylanase 1. Biosci. Biotechnol. Biochem. 72:951-958.[CrossRef][Medline]
    53. 53
  53. Wu, H. C., and M. Tokunaga. 1986. Biogenesis of lipoproteins in bacteria. Curr. Top. Microbiol. Immunol. 125:127-157.[Medline]
    54. 54
  54. Yamaguchi, K., F. Yu, and M. Inouye. 1988. A signal amino acid determinant of the membrane localization of lipoproteins in E. coli. Cell 53:423-432.[CrossRef][Medline]

Journal of Bacteriology, March 2009, p. 1641-1649, Vol. 191, No. 5
0021-9193/09/$08.00+0     doi:10.1128/JB.01285-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.




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