Burkholderia cenocepacia ZmpB Is a Broad-Specificity Zinc Metalloprotease Involved in Virulence
http://www.100md.com
《感染与免疫杂志》
Department of Microbiology and Infectious Diseases, University of Calgary, Calgary, Alberta, Canada T2N 4N1
ABSTRACT
In previous studies we characterized the Burkholderia cenocepacia ZmpA zinc metalloprotease. In this study, we determined that B. cenocepacia has an additional metalloprotease, which we designated ZmpB. The zmpB gene is present in the same species as zmpA and was detected in B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia but was absent from B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina. The zmpB gene was expressed, and ZmpB was purified from Escherichia coli by using the pPROEXHTa His6 Tag expression system. ZmpB has a predicted preproenzyme structure typical of thermolysin-like proteases and is distantly related to Bacillus cereus bacillolysin. ZmpB was expressed as a 63-kDa preproenzyme precursor that was autocatalytically cleaved into mature ZmpB (35 kDa) and a 27-kDa prepropeptide. EDTA, 1,10-phenanthroline, and Zn2+ cations inhibited ZmpB enzyme activity, indicating that it is a metalloprotease. ZmpB had proteolytic activity against -1 proteinase inhibitor, 2-macrogobulin, type IV collagen, fibronectin, lactoferrin, transferrin, and immunoglobulins. B. cenocepacia zmpB and zmpA zmpB mutants had no proteolytic activity against casein and were less virulent in a rat agar bead chronic infection model, indicating that zmpB is involved in B. cenocepacia virulence. Expression of zmpB was regulated by both the CepIR and CciIR quorum-sensing systems.
INTRODUCTION
The Burkholderia cepacia complex is comprised of nine closely related species (8, 9; reviewed in reference 39). B. cepacia complex organisms infect approximately 4 to 7% of cystic fibrosis (CF) patients (12). B. cepacia complex strains may cause a rapid deterioration of lung function and death in some CF patients (27). Strains from all nine species have been isolated from CF patients, but the most prevalent species are B. cenocepacia and B. multivorans, with strains of B. cenocepacia being most commonly isolated from North American CF patients (45, 55). The majority of the transmissible and epidemic B. cepacia complex strains belong to B. cenocepacia (35, 39, 41, 55). Highly transmissible B. cenocepacia strains, such as ET12, Midwest, and PHDC, have been identified in outbreaks in North America and Europe (6, 34). B. cenocepacia ET12 was responsible for the largest B. cepacia complex epidemic affecting CF patients in Canada and the United Kingdom during the late 1980s and early 1990s and has been linked to patient-to-patient transmission (55, 56). B. cenocepacia ET12 strains contain a large number of unique genes (3, 38, 56), although none of these have been directly linked to transmissibility between patients.
Numerous factors that have been implicated in B. cenocepacia virulence have been identified, including protease (11), quorum-sensing systems (1, 30, 54), hemolysin (26), lipopolysaccharide (25), capsule (25), cable pili and 22-kDa adhesin (49), flagella (57), exopolysaccharides (7, 10), siderophores (59), and yfjI, a gene of unknown function (3). We previously identified the zmpA gene and determined that it encodes a zinc metalloprotease that contributes to virulence in chronic lung infections (11). B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia have a zmpA gene and detectable extracellular protease activity, whereas B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina lack zmpA and are protease negative (19). ZmpA has the potential to cause direct tissue damage and to modulate the host immune system, since it has been shown to degrade type IV collagen, fibronectin, -1 proteinase inhibitor, 2-macroglobulin, and gamma interferon (28). ZmpA is expressed as a preproenzyme that is autoproteolytically cleaved into a 36-kDa mature enzyme. It was confirmed to be a zinc metalloprotease, since its activity was inhibited by EDTA and 1,10-phenanthroline (28). Expression of the zmpA gene is regulated by both the CepIR and CciIR quorum-sensing systems (40, 54).
Quorum sensing is a regulatory system that controls expression of target genes in a cell density-dependent manner and in gram-negative bacteria usually involves N-acyl-homoserine lactones (AHLs) as signaling molecules (reviewed in references 44 and 58). AHLs are synthesized by AHL synthases encoded by luxI homologues. The AHLs bind to and activate a response regulator encoded by a luxR homologue that regulates expression of target genes at the level of transcription. Two LuxIR quorum-sensing systems, cepIR and cciIR, have been identified in B. cenocepacia K56-2 (1, 33, 40). CepI and CciI synthesize N-octanoyl-L-homoserine lactone and N-hexanoyl-L-homoserine lactone, although in different ratios (36, 40). The cepIR system is widely distributed throughout the B. cepacia complex (20, 36), whereas the cciIR system is found only in B. cenocepacia ET12 strains that contain the cci genomic island (1).
Protease activity has previously been characterized in B. cenocepacia strains with mutations in zmpA or each of the quorum-sensing genes. B. cenocepacia K56-2 and Pc715j zmpA mutants had significantly less extracellular protease activity than the parent strains; however, the zmpA mutation did not completely eliminate proteolytic activity (11). In fact, the Pc715j zmpA mutant retained approximately 50% of the proteolytic activity of the parent and was equally virulent in the rat agar bead lung infection model (11). B. cenocepacia K56-2 or H111 cepI or cepR mutants produce no detectable protease (24, 32). Interestingly, a K56-2 cciI mutant produced significantly more protease activity than the parent strain, although expression of a zmpA::lacZ fusion was considerably lower in the cciI mutant than the parent strain (40). Taken together, these studies suggested that B. cenocepacia has at least one extracellular protease gene in addition to zmpA and that there are differences between the regulation of zmpA and other protease genes by the cciIR quorum-sensing system. We isolated a spontaneous protease-negative mutant of B. cenocepacia K56-2 that does not express zmpA but has less protease activity than a zmpA mutant, suggesting that it was also deficient in expression of other proteases (data not shown). In the present study, we used this mutant to clone a second metalloprotease gene from B. cenocepacia, which we designated zmpB. We characterized the activity of this protease and demonstrated that it is involved in the virulence of B. cenocepacia and that its expression is regulated by both the CepIR and CciR quorum-sensing systems.
MATERIALS AND METHODS
Strains, plasmids, primers, and growth conditions. Strains and plasmids used in this study are listed in Table 1. Escherichia coli and B. cenocepacia were routinely grown in Luria-Bertani (LB) broth (Invitrogen, Burlington, Ontario, Canada) or on LB agar at 37°C. Bacteria from rat lung homogenates were recovered on Burkholderia cepacia selective agar (21). When appropriate, antibiotics were used at the following concentrations (per milliliter): for B. cenocepacia, 200 μg tetracycline (Tc) and 100 μg trimethoprim (Tp), and for E. coli, 100 μg ampicillin (Ap), 50 μg kanamycin (Km), 15 μg Tc, and 1.5 mg Tp. All chemicals were obtained from Sigma (St. Louis, Mo.) except where indicated.
DNA manipulations. Molecular biology techniques were generally performed as described by Sambrook et al. (50). PCRs were carried out using standard methods as described by the manufacturer, with either Vent DNA polymerase (New England Biolabs, Pickering, Ontario, Canada) for cloning of zmpB or Taq DNA polymerase (Invitrogen) for amplification of a zmpB fragment from B. cepacia complex strains. PCR products or DNA fragments were purified from agarose gels by using a QIAquick gel extraction kit (QIAGEN, Mississauga, Ontario, Canada). DNA sequencing was performed by the University of Calgary Core DNA Sequencing Laboratory. Sequence analysis was performed with DNAMAN software (Lynnon Biosoft, Vaudreuil, Quebec, Canada), and sequences were compared to the unpublished genome sequence of B. cenocepacia J2315 (http://www.sanger.ac.uk/Projects/B_cenocepacia/) by using Artemis software (48).
Cloning of zmpB and construction of zmpB mutants. B. cenocepacia K56-2 genomic DNA was partially digested with Sau3A and cloned into pUCP26 (60) digested with BamHI. The resultant library was electroporated into a spontaneous protease-negative mutant of K56-2 as previously described (14), and transformants were screened for restoration of proteolytic activity on brain heart infusion (BHI)-skim milk agar (53). One positive clone was designated pUCP21D6.
A 2.9-kb PCR fragment containing zmpB was amplified from pUCP21D6 by using primers BS1-21D6F (5'-TGTGCGTCGCGTGCGGCTTC-3') and BS1-21D6R (5'-GTGCGCGACCTGTTCGACGG-3') and cloned into pCR2.1-TOPO (Invitrogen), generating pBS1. For construction of pBS2 and pBS3, antibiotic cassettes (Tpr from p34E-Tp [16] and Tcr from p34S-Tc [15], respectively) were inserted into the pBS1 SalI site after deletion of a 1.4-kb SalI internal fragment.
A 2.1-kb XhoI-HindIII fragment from pBS2 was cloned into the suicide vector pEX18Tc and used to generate K56-2 zmpB by allelic exchange as previously described (22). A 2.9-kb XhoI-HindIII fragment from pBS3 was cloned into pEX18Ap (22) and transformed into B. cenocepacia K56-2-9 (zmpA) (11) to generate K56-2 zmpA zmpB. Allelic exchange mutants were identified by loss of the vector, sucrose resistance, and decreased zones on BHI-skim milk agar. Double-crossover mutants were confirmed by PCR using the primers pBS1-21D6F and BS1-21D6R. For complementation experiments, the 2.9-kb XhoI-HindIII fragment containing zmpB from pBS1 was cloned into pUCP26 (60) to generate pBS6.
Animal studies. The virulence of the K56-2 zmpA, K56-2 zmpB, and K56-2 zmpA zmpB mutants was compared to that of the parent K56-2 by using the agar bead chronic respiratory infection model (4). The number of bacteria and histopathological changes indicating inflammation in the infected lungs were quantified on days 7 and 14 postinfection (p.i.) as previously described (2).
For generation of anti-ZmpB sera, Sprague-Dawley rats (Charles River, Saint Constant, Quebec, Canada) were immunized on days 0 and 21 with recombinant ZmpB (approximately 20 μg) emulsified in monophosphoryl lipid A plus synthetic trehalose dicorynomycolate adjuvant as described by the manufacturer (Corixa Corporation, Hamilton, MT). Rats were bled on day 26 by cardiac puncture and anti-ZmpB antibodies detected by enzyme-linked immunosorbent assay using recombinant ZmpB as the antigen. Production of anti-ZmpA serum was previously described (19).
Expression of the zmpB gene and purification of recombinant ZmpB. The primers ZMPBF (5'-CGGAATTCATGCAGACATCACGCAAAGCATTG) and ZMPBR (5'-CCCAAGCTTCCTGCTGCGGTATTACGACTTCTG) were used to PCR amplify a 1,713-bp fragment containing K56-2 zmpB from the start codon (ATG) to 12 bp beyond the stop codon (TAA). The ZMPBF and ZMPBR primers contain an EcoRI site and a HindIII site (underlined), respectively. The amplified fragment was then digested with EcoRI-HindIII and ligated into pPROEXHTa (Invitrogen) to generate pSCM4.
E. coli DH5(pSCM4) was grown overnight in 10 ml LB supplemented with Ap and used to inoculate 400 ml LB at an initial optical density at 600 nm (OD600) of 0.03. When the cultures reached an OD600 of 0.6, they were induced with 0.7 mM isopropyl--D-thiogalactoside (IPTG) for 3.5 h. Recombinant ZmpB was purified and refolded as previously described for ZmpA (28).
The relative molecular mass and purity of ZmpB were determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (31). The partial N-terminal amino acid sequence of the mature ZmpB was determined from ZmpB separated by SDS-PAGE and electroblotted onto polyvinylidene difluoride (PVDF) membranes (Millipore Corp., Etobicoke, Ontario, Canada). Sequencing was performed by the Protein Microchemistry Laboratory at the University of British Columbia.
To detect ZmpB expression in B. cenocepacia and other B. cepacia complex species, cell-free supernatants were precipitated with trichloroacetic acid to a final concentration of 10% as previously described (11). The proteins were electrophoresed for 30 h at 80 V by Tricine-SDS-14% polyacrylamide gel electrophoresis (51) and transferred to PVDF membranes. ZmpB and ZmpA were detected on the immunoblots with rat antibodies to each protein, using peroxidase-conjugated goat anti-rat immunoglobulin G (IgG) (Kirkegaard & Perry, Gaithersburg, MD).
Protease activity assays. Protease activity was determined on BHI-skim milk agar at 24 and 48 h as previously described (28) or with hide powder azure (47). One unit of protease was defined as the activity that produced a change in the OD595 of 0.1 per hour at 37°C.
The optimal pH for ZmpB proteolytic activity was determined. ZmpB (4 units, 100 ng) was diluted in citrate buffer (25 mM, pH 3 to 6), morpholinoethanesulfonic acid (MES) (25 mM, pH 5 to 6.7), 3-(N-morpholino)propanesulfonic acid (MOPS) (25 mM, pH 6.5 to 8), Tris-HCl (25 mM, pH 8 to 9), or bicarbonate (25 mM, pH 10) and incubated at 37°C for 2 h. The optimum temperature for ZmpB activity was determined in 25 mM MES (pH 5.6). Assays were performed in triplicate with a buffer control at each temperature, using hide powder azure as the substrate.
To identify potential ZmpB substrates, ZmpB (2 units) was incubated for 24 to 48 h at 37°C with -1 proteinase inhibitor (Bayer Corporation, Elkhart, IN), human fibronectin, type IV collagen, lactoferrin, transferrin, IgA, IgG, IgM, rat 2-macroglobulin, and recombinant rat gamma interferon (Sigma) in 25 mM MES, pH 5.6. Recombinant ZmpB and substrate-only controls were included. In addition, substrates were incubated with the proteolytically inactive ZmpA H465G (28) purified in an identical manner as ZmpB to ensure that proteolytic activity was not due to contaminating proteases in the reaction mixtures.
For inhibitor and divalent metal cation studies, ZmpB (4 units, approximately 100 ng) was preincubated with the specified inhibitor(s) and/or divalent metal cation(s) for 30 min at room temperature and transferred to hide powder azure in 25 mM MES (pH 5.6). Neutralization assays with a monoclonal antibody (36-6-8) to ZmpA were performed as previously described (29).
Expression of zmpB::luxCDABE transcriptional fusions. A zmpB::luxCDABE transcriptional fusion was constructed in pMS402 (17). A 0.75-kb DNA fragment containing the zmpB promoter was PCR amplified from K56-2 genomic DNA with BSpzmpBF (5'-ATACTTTCGTATCGGCCAC-3') and BSpzmpBR (5'-ACGCACTTCTACCTGAACCT-3'). The amplified fragment was digested with BamHI-XhoI and ligated into the BamHI-XhoI site upstream of luxCDABE in pMS402, resulting in pBS8. To generate pBS9, a SacI fragment from p34STc (15) containing the tetracycline resistance cassette was ligated into pBS8 digested with XhoI.
Luminescence assays were performed to measure expression of zmpB. Overnight cultures were subcultured in triplicate to an initial OD600 of 0.02 in 20 ml LB and grown at 37°C and 250 rpm. At 16 and 20 h, 150-μl aliquots were transferred into 96-well clear-bottom black plates (9520 Costar; Corning Incorporated, Corning, NY), and the luminescence in counts per second and turbidity at an absorbance of 600 nm were measured using a Wallac Victor2 1420 multilabel counter (Perkin-Elmer Life Sciences, Boston, Mass.). Expression levels of zmpB were determined by relative luminescence.
Distribution of the zmpB gene in the B. cepacia complex. To determine the distribution of the zmpB gene in Burkholderia species, primers ZPBDTF (GCAGACATCACGCAAAGCATTG) and ZPBDTR (GCAGCTCTTGTTGTACGCGTTG) were used to PCR amplify a 1.7-kb zmpB fragment. The amplification conditions were 95°C for 7 min; followed by 30 cycles of 95°C for 1 min, 56°C for 30 s, and 72°C for 2 min; followed by 72°C for 20 min.
Southern hybridization analysis was also performed on genomic DNA isolated from representative strains digested with NotI. A 0.8-kb MluI fragment encoding from Arg-35 to Asn-307 of the 323-amino-acid mature ZmpB was labeled with 32P by random priming as described by the manufacturer (GE Healthcare) and used as a probe. Blots were hybridized with the probe at 65°C and washed at 72°C.
RESULTS
Identification of the zmpB gene. The plasmid pUCP21D6 with a 6,780-bp insert containing a putative metalloprotease gene was identified by complementation of a spontaneous protease-negative mutant that does not express zmpA (data not shown) with a B. cenocepacia K56-2 genomic library. Analysis of this insert by using the unpublished annotation of B. cenocepacia J2315 at the Sanger Institute (http://www.sanger.ac.uk/Projects/B_cenocepacia/) and Artemis software (48) revealed five open reading frames encoding two transcriptional regulators of the AraC family (BCAM2305 and BCAM2306), a thermolysin metalloprotease (BCAM2307), a leucyl aminopeptidase (BCAM2308), and a hypothetical protein (BCAM2309). BCAM2307, encoding a putative thermolysin-like metalloprotease belonging to the M4 family of peptidases, was designated zmpB, and its corresponding protein was designated ZmpB. Alignment analysis of ZmpB with the proteases most related to it, bacillolysin from Bacillus cereus ATCC 14579 (29% identity) and thermolysin peptidase M4 from Bacillus anthracis A2012 (27% identity), indicated that the HExxH active-site motif and amino acids downstream of the active site (including GxxxExxxD, where E is a putative zinc ligand) are conserved. Functional domain analysis using the Conserved Domain Database (http://www.ncbi.nlm.nih.gov/Structure/cdd) demonstrated that ZmpB is also functionally related to Pseudomonas aeruginosa LasB and has a prepropeptide structure typical of secreted metalloproteases. The precursor to ZmpB has a theoretical pI of 5.84 and a molecular mass of 60547 Da (http://us.expasy.org.). The pre sequence has a hydrophobic transmembrane domain at amino acids 7 to 26 (http://www.sbc.su.se/miklos/DAS) and a predicted SignalP recognition cleavage site 24TVA. The N-terminal amino acid sequence (FVFRPDPLS) was determined with a recombinant ZmpB peptide described below. Based on this amino acid sequence, the mature ZmpB has a predicted molecular mass of 34.9 kDa and pI of 5.08.
Characterization of K56-2 zmpB and K56-2 zmpA zmpB mutants. Mutation of the zmpB gene in K56-2 and K56-2-9 zmpA was performed by insertional inactivation of zmpB with a Tpr or Tcr cassette, resulting in K56-2 zmpB and K56-2 zmpA zmpB, respectively. The extracellular protein profiles were analyzed by Tricine-SDS-14% PAGE (Fig. 1). K56-2 and K56-2-9 zmpA expressed a 35-kDa protein, which corresponded to the predicted molecular mass of ZmpB. The 35-kDa protein was not detected in K56-2 zmpB and K56-2 zmpA zmpB supernatants (Fig. 1A). The 35-kDa proteins of K56-2, K56-2-9 zmpA, and K56-2 zmpB(pBS6) reacted with anti-rat ZmpB antibody (Fig. 1B), indicating that pBS6 complemented K56-2 zmpB. The 36-kDa peptides present in K56-2, K56-2 zmpB, and K56-2 zmpB (pBS6) correspond to the molecular mass of ZmpA (29, 42) and reacted with anti-ZmpA (Fig. 1C). Anti-ZmpA and anti-ZmpB antibodies did not cross-react with the heterologous proteases, suggesting that these two proteases are antigenically distinct.
FIG. 1. Expression of ZmpA and ZmpB in K56-2 zmpB and K56-2 zmpA zmpB mutants. Supernatant proteins were precipitated with 10% trichloroacetic acid, separated by Tricine-14% SDS-PAGE, and blotted to PVDF membranes. A. Coomassie blue-stained gel. B. Western blot reacted with anti-ZmpB. C. Western blot reacted with anti-ZmpA. Lanes 1, K56-2; lanes 2, K56-2 zmpB; lanes 3, K56-2 zmpB(pBS6); lanes 4, K56-2 zmpA zmpB; lanes 5, K56-2 zmpA; lanes 6, recombinant ZmpB; lanes 7, recombinant ZmpA. Solid arrow, ZmpA; open arrow, ZmpB.
The proteolytic activities of K56-2, K56-2 zmpB, and K56-2 zmpA zmpB mutants were determined on BHI-skim milk agar (Table 2). K56-2 had significantly more protease activity than K56-2 zmpB and K56-2 zmpA zmpB. K56-2 zmpB(pBS6) had zones comparable in size to those of K56-2, indicating that pBS6, containing zmpB, complemented the protease-negative phenotype of K56-2 zmpB. The protease activity of K56-2-9 zmpA was similar to that previously reported (11). These results indicate that zmpB is responsible for the majority of the protease activity of B. cenocepacia K56-2. Complementation of K56-2 zmpB with pBS6 did not result in increased protease activity above that of the wild type despite the potential for increased amounts of ZmpB protein due to the multicopy plasmid. Similar results were reported for complementation of zmpA mutants (11).
The effect of a zmpB mutation on virulence was examined using a chronic respiratory infection model. Previously, we reported that the K56-2-9 zmpA mutant was less persistent in lungs and manifested less histopathology than the parent K56-2 (11). Similar results were obtained in this study in that the number of K56-2-9 zmpA or K56-2 zmpA zmpB organisms recovered from the lungs at day 14 p.i. was approximately three log units lower than the number of K56-2 organisms recovered (Table 3). Three of the four rats infected with K56-2-9 zmpA were completely cleared of bacteria by day 14 p.i. Bacteria recovered from the fourth rat were determined by sequence analysis to be revertants. The numbers of K56-2 zmpB organisms recovered from the lungs were similar to those of K56-2 at both 7 and 14 days p.i. (Table 3), suggesting that zmpA plays a role in bacterial persistence but that zmpB does not.
Both zmpA and zmpB contribute to lung pathology. The lungs from the rats infected with the K56-2 zmpA, K56-2 zmpB, or K56-2 zmpA zmpB protease mutant had significantly fewer histopathological changes than the lungs from rats infected with the K56-2 parent (Table 3), indicating that both proteases contribute to the maximal virulence of B. cenocepacia K56-2 in this infection model. There was no significant difference in the quantitative histopathological changes in the K56-2 zmpA zmpB double mutant and the zmpA or zmpB single mutant.
Purification and characteristics of recombinant ZmpB. The zmpB gene was cloned into pPROEXHTa to construct pSCM4 and was expressed as an N-terminal His-tagged protein. E. coli DH5(pSCM4) was proteolytically active on BHI-skim milk agar, with zones of approximately 2 to 4 mm after 48 h, in contrast to E. coli DH5(pPROEXHTa), which displayed no proteolytic activity. The zmpB gene was expressed as a 63-kDa protein in the cell lysate after IPTG induction for 3.5 h (Fig. 2, lane 1). The mass of 63 kDa corresponded to the predicted molecular mass of the His6-tagged ZmpB (63.4 kDa). When the cell lysate was fractionated and analyzed by SDS-PAGE, the 63-kDa protein was recovered almost exclusively in the precipitate as insoluble inclusion bodies (Fig. 2, lane 3). When the inclusion bodies were solubilized in 6 M guanidine-HCl and applied to an Ni-nitrilotriacetic acid (NTA) affinity column to purify and refold the 63-kDa protein, two new peptides (35 kDa and 28 kDa) were identified in the eluant (Fig. 2, lane 4). The N-terminal sequence of the 35-kDa peptide was determined to be FVFRPDPLS, which is present in the C-terminal portion of the 63-kDa His6-tagged ZmpB, and this mass corresponds to the predicted mass of 34.9 kDa that would be obtained for a peptide with this N-terminal sequence. Therefore, the 35-kDa peptide contains the active site and is the mature protease. The 28-kDa peptide has the predicted mass for the prepropeptide and the His tag.
FIG. 2. Expression and purification of ZmpB. Lane M, molecular weight markers (in thousands); lane 1, cell lysate after induction with IPTG; lane 2, uninduced crude cell extract; lane 3, purified inclusion bodies; lane 4, purified recombinant ZmpB after refolding and elution from an Ni-NTA column. Solid arrow, His-prepro-ZmpB fusion protein; open arrow, indicates the His-prepropeptide. The mature ZmpB is indicated.
ZmpB eluted from the Ni-NTA column was proteolytically active against hide powder azure, with 50 ng containing approximately 5 U of activity. The effect of protease inhibitors or divalent cations on ZmpB protease activity was examined. ZmpB activity was inhibited by EDTA and 1,10-phenanthroline, which chelate Zn2+, but not by phenylmethylsulfonyl fluoride or phosphoramidon (Table 4). ZmpB was not inhibited by 1,7-phenanthroline, an isomer of 1,10-phenanthroline that does not bind Zn2+. In addition, 1 mM Zn2+ abrogated ZmpB activity, but 1 mM CaCl2 had no effect (Table 4). Co2+ and Ni2+ also significantly reduced ZmpB activity. Monoclonal antibody 36-6-8, which previously was shown to inhibit ZmpA, did not inhibit ZmpB activity (29). The A595 when ZmpB was preincubated with monoclonal antibody 36-6-8 was 0.7 ± 0.2, compared with 0.7 ± 0.1 in control ascites fluid.
The optimal pH range for ZmpB activity was between pH 5.6 and 7.2, depending on the buffers employed (data not shown). ZmpB was active over a broad pH range, although activity was dramatically decreased at pH values below 4.5 or above 8.0. ZmpB was active between 28 and 60°C, with an optimum temperature of 50°C (data not shown). Activity was significantly decreased at temperatures above 70°C.
ZmpB activity against host proteins. The ability of ZmpB to degrade a number of biologically important substrates which might account for its role in virulence was determined by incubating ZmpB with the substrate and determining proteolysis by SDS-PAGE (Fig. 3). ZmpB was able to digest the human plasma protease inhibitors -1 proteinase inhibitor and 2-macroglobulin. ZmpB also digested type IV collagen, fibronectin, transferrin, lactoferrin, and immunoglobulins. Collagen, fibronectin, -1 proteinase inhibitor, and 2-macroglobulin were cleaved into numerous peptides, indicating that ZmpB has broad specificity. Transferrin, lactoferrin, and the immunoglobulins IgA, IgG, and IgM were also cleaved into several peptides but were less extensively degraded by ZmpB than collagen, fibronectin, -1 proteinase inhibitor, and 2-macroglobulin.
FIG. 3. Substrate specificity of ZmpB. Substrates were digested with ZmpB, ZmpAH465G (which has no proteolytic activity), or in the absence of enzyme (no protease) for 24 to 48 h. Proteins were separated by SDS-PAGE and stained with Coomassie blue. Collagen, fibronectin, and 2-macroglobulin were electrophoresed by SDS-7.5% PAGE. All other substrates were electrophoresed by SDS-12.5% PAGE.
In order to ensure that the ZmpB activity was not due to the presence of a contaminating E. coli protease, these substrates were digested with a recombinant inactive form of ZmpA, ZmpA H465G (28), that was purified using the same protocol. ZmpA H465G had no activity against any of the substrates, confirming that the cleavage of all substrates was not due to the presence of contaminating proteases in the reaction mixtures or proteases that might copurify from E. coli.
Quorum-sensing regulation of zmpB expression. Previously, zmpA had been shown to be regulated by the cepIR and cciIR quorum-sensing systems (40, 54). To determine if zmpB expression was regulated by cepIR or cciIR, the zmpB promoter region was cloned upstream of the luxCDABE operon in pMS402 (17). The resulting plasmid, pBS9, was transformed into K56-2, K56-dI2 cepI, K56-R2 cepR, K56-2 cciI, K56-2 cciR, K56 cciIR, and K56-2 cepR cciIR. Expression of zmpB::luxCDABE was reduced to almost background levels in K56-dI2(pBS9) and K56-R2(pBS9) and was significantly reduced in K56-2 cciR(pBS9) and K56-2 cepR cciIR(pBS9) (Fig. 4). Interestingly, zmpB::luxCDABE was significantly increased in K56-2 cciI(pBS9). Expression in K56-2 cciIR(pBS9) was similar to that in the parent. These data suggest that zmpB expression is regulated by both cepIR and cciIR.
FIG. 4. Regulation of zmpB by the cepIR and cciIR quorum-sensing genes. Plasmid pBS9 containing a zmpB::luxCDABE fusion was introduced into K56-2 quorum-sensing mutants. Cultures were grown for 20 h, and luminescence in counts per second (CPS) and turbidity at an absorbance of 600 nm were measured using a Wallac Victor2 1420 multilabel counter. Values shown are the mean relative CPS (±standard deviation) for triplicate cultures and are representative of three independent experiments. , significantly different from result for K56-2 (P < 0.05 by analysis of variance).
Distribution of the zmpB gene in Burkholderia spp. To investigate the distribution of the zmpB gene in Burkholderia species, primers ZPBDTF and ZPBDTR were used to PCR amplify the 1.7-kb zmpB gene in 43 strains representing the B. cepacia complex. A 1.7-kb amplicon was obtained from all strains of B. cepacia, B. cenocepacia, B. ambifaria, and B. pyrrocinia examined but not from strains of B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina (data not shown). Southern blot hybridization analysis was also performed to determine which species contained a zmpB gene (Fig. 5A). B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia had a 0.8-kb NotI band that hybridized to the zmpB probe as strongly as that of the positive control strain K56-2 (Fig. 5A). The probe did not hybridize to B. multivorans, B. vietnamiensis, B. dolosa, or B. anthina, suggesting that these species do not contain a gene related to zmpB, although it is possible that these species could contain a related gene with less homology than would be detected by Southern hybridization.
FIG. 5. Distribution of ZmpB in the B. cepacia complex. A. Southern hybridization analysis of the zmpB gene. A 0.8-kb 32P-labeled MluI fragment encoding from Arg-35 to Asn-307 of the 323-amino-acid mature ZmpB from K56-2 was used to probe NotI-digested genomic DNA. Lane 1, K56-2; lane 2, B. cepacia LMG17997; lane 3, B. multivorans LMG13010; lane 4, B. cenocepacia J2315; lane 5, B. stabilis LMG14294; lane 6, B. stabilis C7322; lane 7, B. vietnamiensis PC259; lane 8, B. dolosa LMG18943; lane 9, B. ambifaria Cep996; lane 10, B. anthina LMG20980; lane 11, B. pyrrocinia LMG21822. B. Detection of ZmpB by Western blotting of culture supernatants. Trichloroacetic acid-precipitated culture supernatants were separated by Tricine 14%-PAGE and blotted onto PVDF membrane. Blots were reacted with anti-ZmpB. Lane 1, K56-2; lane 2, B. cepacia LMG17997; lane 3, B. multivorans LMG13010; lane 4, B. cenocepacia C6433; lane 5, B. stabilis LMG18888; lane 6, B. vietnamiensis Pc259; lane 7, B. dolosa LMG18943; lane 8, B. ambifaria CEP996; lane 9, B. anthina LMG20980; lane 10, B. pyrrocinia LMG 21822; lane 11, recombinant ZmpB.
To determine if the presence of zmpB correlated with extracellular protease activity, we examined the activity of 43 strains on BHI-skim milk agar. The majority (18 out of 23) of the zmpB-positive strains had extracellular protease activity, while none of the zmpB-negative strains produced detectable extracellular protease activity. These data suggest that the strains in which zmpB was not detectable by PCR or Southern hybridization do not contain a related gene. ZmpB was easily detected only in B. cenocepacia concentrated culture supernatants by Western blot analysis (Fig. 5B). Possibly ZmpB was not expressed and/or secreted in the other zmpB-positive B. cepacia complex strains under the conditions employed, or ZmpB was secreted but the anti-ZmpB polyclonal antibody employed did not react with the ZmpB from these strains.
DISCUSSION
Metalloproteases are widely distributed in nature and belong to four superfamilies or clans. All bacterial zinc metalloproteases belong to the zincins superfamily, which contains a conserved HEXXH motif. The two histidines are zinc ligands, and the glutamic acid is involved in enzymatic activity. The zincins can be subdivided, based on the position of the third zinc ligand (43), into at least 10 families, with 3 of the families containing bacterial metalloproteases: thermolysin, serralysin, and neurotoxins.
B. cenocepacia ZmpB is a novel thermolysin-like protease that has only 28% identity with its closest homologue, bacillolysin from Bacillus cereus, and shares only 10.3% identity with B. cenocepacia ZmpA. ZmpB has a preproenzyme structure that is typical of the M4 family (http://merops.sanger.ac.uk/) of metalloproteases, with conservation of functional regions such as the HExxH active-site motif as well as other conserved amino acids downstream of the active site, including GxxxExxxD. De Kreij et al. (13) found that there was a great deal of sequence variability among various thermolysin-like proteases but that as long as a small subset of amino acids was conserved (e.g., active site residues and zinc ligands), the group was extremely homogeneous in terms of catalysis. B. cenocepacia ZmpB is inhibited by EDTA and 1,10-phenanthroline, indicating that it is a metalloprotease. ZmpB is thermostable and active over a broad neutral pH range, similar to other bacterial thermolysin-like metalloproteases. ZmpB activity was relatively unaffected by Ca2+ and Mg2+ but completely abrogated by excess Zn2+, as has been reported with thermolysin (23). Co2+ and to lesser degree Mn2+ could substitute for Zn2+ in thermolysin (23), which is also suggested by our findings for ZmpB. Ni2+ has not been shown to inhibit thermolysin, but Ni2+, Zn2+, and Cu2+ have been shown to inhibit a Vibrio vulnificus broad-specificity metalloprotease (5). Interestingly, ZmpB was not inhibited by phosphoramidon, a peptide inhibitor of metalloproteases that is related to B. thermoproteolyticus thermolysin. B. cenocepacia ZmpA is also unaffected by phosphoramidon (28). Phosphoramidon is a peptide that binds to a specific sequence in the thermolysin active-site region. Sequence differences in this region have been correlated with reduced phosphoramidon sensitivity (13).
ZmpB was shown to be a broad-spectrum protease capable of degrading substrates involved in tissue integrity and host defense. Most proteases are inhibited by 2-macroglobulin, but both ZmpB and ZmpA (28) are able to digest 2-macroglobulin and -1 proteinase inhibitor, indicating that B. cenocepacia produces two extracellular proteases that can inactivate the major mammalian protease inhibitors. Inactivation of -1 proteinase inhibitor in vivo may disrupt the -1 proteinase/-1 proteinase inhibitor balance and lead to increased tissue damage and inflammation. Inactivation of 2-macroglobulin could result in increased B. cenocepacia dissemination and lead to septicemia. Both ZmpB and ZmpA can degrade host tissue components, including collagen and fibronectin. Although ZmpB and ZmpA have activity against many of the same substrates, each enzyme has a different specificity. ZmpA cleaved -1 proteinase inhibitor specifically at D36 (D36TSHHDQD) (28), whereas ZmpB cleaved -1 proteinase inhibitor into a number of fragments (Fig. 3). ZmpA specifically cleaved 2-macroglobulin into major polypeptides of 95 and 86 kDa (28). ZmpB cleaved 2-macroglobulin into numerous peptides (Fig. 3).
ZmpB was shown to cleave immunoglobulins (IgA, IgG, and IgM), transferrin, and lactoferrin, which could also increase the ability of B. cenocepacia to evade host defenses. Although we previously cited unpublished data (52) that suggested that ZmpA had activity against these substrates, recombinant and native ZmpA were subsequently shown not to degrade these substrates (28). It is likely that the protease preparations used in the unpublished study contained a mixture of both ZmpA and ZmpB proteases. ZmpB digested lactoferrin, transferrin, and immunoglobulins into markedly fewer peptides than 2-macroglobulin, collagen, and fibronectin, suggesting that there is some cleavage site specificity. ZmpA was shown to digest gamma interferon (28). Because ZmpB had significant proteolytic activity against the albumin present in large amounts in the gamma interferon preparation used, it was not possible to conclusively distinguish between gamma interferon and albumin cleavage products in this assay.
ZmpA was previously shown to be necessary for B. cenocepacia K56-2 to cause persistent respiratory infections in rats (11). Similar results were obtained in the present study, in that the numbers of K56-2 zmpA and K56-2 zmpA zmpB organisms recovered from infected lungs were approximately 3 log units lower than in the parent strain. In contrast, K56-2 zmpB was able to survive at parental levels in the rat lung, indicating that ZmpB is not required for persistence in this strain. Infection with any of the mutants resulted in significantly less lung inflammation than infection with the parent strain. There was no significant difference between the quantitative histopathology changes in lungs infected with the mutants. The decreased inflammation observed in lungs infected with zmpA mutants compared to the parent is at least partially due to the decreased numbers of bacteria present in the lung, whereas the decreased inflammation observed with the zmpB mutant is likely due to the absence of this enzyme. Although a zmpA mutation in K56-2 resulted in decreased virulence, a Pc715j zmpA mutant was virulent in the lung infection model (11). Unfortunately, we were not able to construct a zmpB mutation in either Pc715j or Pc715j zmpA to determine its effects on virulence.
Although zmpA and zmpB are not genetically linked and are in fact located on different chromosomes, they are present in the same B. cepacia complex species. Both genes are present in B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia but are absent from B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina (19). Presence or absence of these genes is conserved within each species. Highly (>95%) conserved ZmpB homologues are present in the genomes of B. cepacia R18194, B. cenocepacia J2315, B. cenocepacia HI2424, and B. ambifaria AMMD.
Some B. cepacia complex strains were previously reported to have no detectable protease activity despite having the zmpA gene (19). Others had similar levels of protease activity, but the induction of protease production appeared to be much slower. Interestingly, all of these zmpA-positive, protease-deficient strains are also zmpB positive, suggesting that zmpB may also be poorly expressed or nonfunctional in these strains. These data suggest that zmpA and zmpB share similar regulatory mechanisms. It is also possible that these enzymes act synergistically, since they have different cleavage properties. The zmpB mutant has no activity against casein in the BHI-skim milk agar assay, and the zmpA mutant has approximately 20% of the activity of the parent, suggesting that these enzymes may work together to degrade casein, resulting in zones of clearance. The histopathologies observed in lungs of infected rats were similar for animals infected with either the zmpA, zmpB, or zmpA zmpB double mutant, again suggesting that the activities of these enzymes are not additive, since knocking out either gene results in a similar reduction in virulence.
Previously, we have reported that mutations in cepI, cepR, cciI, or cciR result in decreased zmpA expression in K56-2 (40, 54). In this study, we determined that zmpB expression was also markedly reduced in cepI, cepR, cciR, and cepR cciIR mutants. The expression of the zmpB::luxCDABE fusion was not above the background fluorescence obtained with the pMS402 vector control in the cepI or cepR mutants, indicating that the cepIR quorum-sensing genes are required for expression of both zmpB and zmpA. These results correlate with the observed absence of protease activity in cepI and cepR mutants (32, 40). The cciR and cepR cciR mutants were also previously shown to have decreased protease activity compared to the parent strain, which correlates with the zmpB expression data obtained in this study. Interestingly, the cciI mutant expressed zmpB::luxCDABE at much higher levels than the parent or any of the other mutants. The increase in zmpB expression corresponds to the increased protease activity previously observed in the cciI mutant on skim milk agar (40). The zones of protease activity in the cciI mutant could be restored to parental levels by the addition of 2.5 nM N-hexanoyl-L-homoserine lactone, which is the major AHL synthesized by cciI. These data suggest that the protease activity observed in the cciI mutant is due to the increased expression of zmpB and that the observed decrease in zmpA expression has little impact on total protease activity. It is interesting that the presence of zmpB on a multicopy plasmid does not result in increased protease zone sizes similar to that observed in the cciI mutant but only restores protease activity to the parental level. This finding suggests that zmpB copy number is not sufficient for increased protease expression or activity but that other factors, such as the balance of AHL signals, have a significant influence on the amount of protease produced. Mutations in cciI and cciR result in different effects on zmpB expression and protease activity. These mutants also have different swarming phenotypes, suggesting that other genes must also be differentially expressed in cciI and cciR mutants (40).
The ZmpB precursor is processed into mature ZmpB and a propeptide with a predicted molecular mass of 23.1 kDa and a pI of 8.8. The propeptide mass and pI correspond to a peptide identified in a proteome comparison between B. cenocepacia H111 and an H111 cepI mutant (46). The QS8 ID spot was identified as an intracellular thermolysin metallopeptidase and presumed to be a degradation product of a precursor metalloprotease. The N-terminal sequence of QS8 was determined to be DTKVSN, which corresponds to the predicted N terminus of the ZmpB propeptide after removal of the pre-signal peptide (46). Peptide QS8 was found to be approximately 20-fold up-regulated in the H111 parent compared to the cepI mutant, which is consistent with our data indicating that zmpB is positively regulated by the cepIR system.
In summary, we have shown that B. cenocepacia has two zinc metalloproteases that contribute to virulence and have the potential to interfere with host defense mechanisms. Further studies are under way in our laboratory to determine the in vivo activities of these proteases and their mechanisms of regulation.
ACKNOWLEDGMENTS
This work was supported by grants from the Canadian Institutes of Health Research and the Canadian Cystic Fibrosis Foundation to P.A.S.
We thank members of the Sokol laboratory for helpful discussions and critical review of the manuscript.
FOOTNOTES
These authors contributed equally to this study.
REFERENCES
1. Baldwin, A., P. A. Sokol, J. Parkhill, and E. Mahenthiralingam. 2004. The Burkholderia cepacia epidemic strain marker is part of a novel genomic island encoding both virulence and metabolism-associated genes in Burkholderia cenocepacia. Infect. Immun. 72:1537-1547.
2. Bernier, S. P., L. Silo-Suh, D. E. Woods, D. E. Ohman, and P. A. Sokol. 2003. Comparative analysis of plant and animal models for characterization of Burkholderia cepacia virulence. Infect. Immun. 71:5306-5313.
3. Bernier, S. P., and P. A. Sokol. 2005. Use of suppression-subtractive hybridization to identify genes in the Burkholderia cepacia complex that are unique to Burkholderia cenocepacia. J. Bacteriol. 187:5278-5291.
4. Cash, H. A., D. E. Woods, B. McCullough, W. G. Johanson, Jr., and J. A. Bass. 1979. A rat model of chronic respiratory infection with Pseudomonas aeruginosa. Am. Rev. Respir. Dis. 119:453-459.
5. Chang, A. K., H. Y. Kim, J. E. Park, P. Acharya, I. S. Park, S. M. Yoon, H. J. You, K. S. Hahm, J. K. Park, and J. S. Lee. 2005. Vibrio vulnificus secretes a broad-specificity metalloprotease capable of interfering with blood homeostasis through prothrombin activation and fibrinolysis. J. Bacteriol. 187:6909-6916.
6. Chen, J. S., K. A. Witzmann, T. Spilker, R. J. Fink, and J. J. LiPuma. 2001. Endemicity and inter-city spread of Burkholderia cepacia genomovar III in cystic fibrosis. J. Pediatr. 139:643-649.
7. Chung, J. W., E. Altman, T. J. Beveridge, and D. P. Speert. 2003. Colonial morphology of Burkholderia cepacia complex genomovar III: implications in exopolysaccharide production, pilus expression, and persistence in the mouse. Infect. Immun. 71:904-909.
8. Coenye, T., P. Vandamme, J. R. Govan, and J. J. LiPuma. 2001. Taxonomy and identification of the Burkholderia cepacia complex. J. Clin. Microbiol. 39:3427-3436.
9. Coenye, T., P. Vandamme, J. J. LiPuma, J. R. Govan, and E. Mahenthiralingam. 2003. Updated version of the Burkholderia cepacia complex experimental strain panel. J. Clin. Microbiol. 41:2797-2798.
10. Conway, B. A., K. K. Chu, J. Bylund, E. Altman, and D. P. Speert. 2004. Production of exopolysaccharide by Burkholderia cenocepacia results in altered cell-surface interactions and altered bacterial clearance in mice. J. Infect. Dis. 190:957-966.
11. Corbett, C. R., M. N. Burtnick, C. Kooi, D. E. Woods, and P. A. Sokol. 2003. An extracellular zinc metalloprotease gene of Burkholderia cepacia. Microbiology 149:2263-2271.
12. Cystic Fibrosis Foundation. 2002. Cystic Fibrosis Foundation patient registry annual data report. Cystic Fibrosis Foundation, Bethesda, Md.
13. de Kreij, A., G. Venema, and B. van den Burg. 2000. Substrate specificity in the highly heterogeneous M4 peptidase family is determined by a small subset of amino acids. J. Biol. Chem. 275:31115-31120.
14. Dennis, J. J., and P. A. Sokol. 1995. Electrotransformation of Pseudomonas. Methods Mol. Biol. 47:125-133.
15. Dennis, J. J., and G. J. Zylstra. 1998. Plasposons: modular self-cloning minitransposon derivatives for rapid genetic analysis of gram-negative bacterial genomes. Appl. Environ. Microbiol. 64:2710-2715.
16. DeShazer, D., and D. E. Woods. 1996. Broad-host-range cloning and cassette vectors based on the R388 trimethoprim resistance gene. BioTechniques 20:762-764.
17. Duan, K., C. Dammel, J. Stein, H. Rabin, and M. G. Surette. 2003. Modulation of Pseudomonas aeruginosa gene expression by host microflora through interspecies communication. Mol. Microbiol. 50:1477-1491.
18. Figurski, D. H., and D. R. Helinski. 1979. Replication of an origin-containing derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl. Acad. Sci. USA 76:1648-1652.
19. Gingues, S., C. Kooi, M. B. Visser, B. Subsin, and P. A. Sokol. 2005. Distribution and expression of the ZmpA metalloprotease in the Burkholderia cepacia complex. J. Bacteriol. 187:8247-8255.
20. Gotschlich, A., B. Huber, O. Geisenberger, A. Togl, A. Steidle, K. Riedel, P. Hill, B. Tummler, P. Vandamme, B. Middleton, M. Camara, P. Williams, A. Hardman, and L. Eberl. 2001. Synthesis of multiple N-acylhomoserine lactones is wide-spread among the members of the Burkholderia cepacia complex. Syst. Appl. Microbiol. 24:1-14.
21. Henry, D., M. Campbell, C. McGimpsey, A. Clarke, L. Louden, J. L. Burns, M. H. Roe, P. Vandamme, and D. Speert. 1999. Comparison of isolation media for recovery of Burkholderia cepacia complex from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 37:1004-1007.
22. Hoang, T. T., R. R. Karkhoff-Schweizer, A. J. Kutchma, and H. P. Schweizer. 1998. A broad-host-range Flp-FRT recombination system for site-specific excision of chromosomally-located DNA sequences: application for isolation of unmarked Pseudomonas aeruginosa mutants. Gene 212:77-86.
23. Holland, D. R., A. C. Hausrath, D. Juers, and B. W. Matthews. 1995. Structural analysis of zinc substitutions in the active site of thermolysin. Protein Sci. 4:1955-1965.
24. Huber, B., K. Riedel, M. Hentzer, A. Heydorn, A. Gotschlich, M. Givskov, S. Molin, and L. Eberl. 2001. The cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility. Microbiology 147:2517-2528.
25. Hunt, T. A., C. Kooi, P. A. Sokol, and M. A. Valvano. 2004. Identification of Burkholderia cenocepacia genes required for bacterial survival in vivo. Infect. Immun. 72:4010-4022.
26. Hutchison, M. L., I. R. Poxton, and J. R. Govan. 1998. Burkholderia cepacia produces a hemolysin that is capable of inducing apoptosis and degranulation of mammalian phagocytes. Infect. Immun. 66:2033-2039.
27. Isles, A., I. Maclusky, M. Corey, R. Gold, C. Prober, P. Fleming, and H. Levison. 1984. Pseudomonas cepacia infection in cystic fibrosis: an emerging problem. J. Pediatr. 104:206-210.
28. Kooi, C., C. R. Corbett, and P. A. Sokol. 2005. Functional analysis of the Burkholderia cenocepacia ZmpA metalloprotease. J. Bacteriol. 187:4421-4429.
29. Kooi, C., A. Cox, P. Darling, and P. A. Sokol. 1994. Neutralizing monoclonal antibodies to an extracellular Pseudomonas cepacia protease. Infect. Immun. 62:2811-2817.
30. Kothe, M., M. Antl, B. Huber, K. Stoecker, D. Ebrecht, I. Steinmetz, and L. Eberl. 2003. Killing of Caenorhabditis elegans by Burkholderia cepacia is controlled by the cep quorum-sensing system. Cell. Microbiol. 5:343-351.
31. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.
32. Lewenza, S., B. Conway, E. P. Greenberg, and P. A. Sokol. 1999. Quorum sensing in Burkholderia cepacia: identification of the LuxRI homologs CepRI. J. Bacteriol. 181:748-756.
33. Lewenza, S., and P. A. Sokol. 2001. Regulation of ornibactin biosynthesis and N-acyl-L-homoserine lactone production by CepR in Burkholderia cepacia. J. Bacteriol. 183:2212-2218.
34. Lipuma, J. J. 2005. Update on the Burkholderia cepacia complex. Curr. Opin. Pulm. Med. 11:528-533.
35. LiPuma, J. J., T. Spilker, L. H. Gill, P. W. Campbell III, L. Liu, and E. Mahenthiralingam. 2001. Disproportionate distribution of Burkholderia cepacia complex species and transmissibility markers in cystic fibrosis. Am. J. Respir. Crit. Care Med. 164:92-96.
36. Lutter, E., S. Lewenza, J. J. Dennis, M. B. Visser, and P. A. Sokol. 2001. Distribution of quorum-sensing genes in the Burkholderia cepacia complex. Infect. Immun. 69:4661-4666.
37. Mahenthiralingam, E., T. Coenye, J. W. Chung, D. P. Speert, J. R. Govan, P. Taylor, and P. Vandamme. 2000. Diagnostically and experimentally useful panel of strains from the Burkholderia cepacia complex. J. Clin. Microbiol. 38:910-913.
38. Mahenthiralingam, E., D. A. Simpson, and D. P. Speert. 1997. Identification and characterization of a novel DNA marker associated with epidemic Burkholderia cepacia strains recovered from patients with cystic fibrosis. J. Clin. Microbiol. 35:808-816.
39. Mahenthiralingam, E., T. A. Urban, and J. B. Goldberg. 2005. The multifarious, multireplicon Burkholderia cepacia complex. Nat. Rev. Microbiol. 3:144-156.
40. Malott, R. J., A. Baldwin, E. Mahenthiralingam, and P. A. Sokol. 2005. Characterization of the cciIR quorum-sensing system in Burkholderia cenocepacia. Infect. Immun. 73:4982-4992.
41. McDowell, A., E. Mahenthiralingam, J. E. Moore, K. E. Dunbar, A. K. Webb, M. E. Dodd, S. L. Martin, B. C. Millar, C. J. Scott, M. Crowe, and J. S. Elborn. 2001. PCR-based detection and identification of Burkholderia cepacia complex pathogens in sputum from cystic fibrosis patients. J. Clin. Microbiol. 39:4247-4255.
42. McKevitt, A. I., S. Bajaksouzian, J. D. Klinger, and D. E. Woods. 1989. Purification and characterization of an extracellular protease from Pseudomonas cepacia. Infect. Immun. 57:771-778.
43. Miyoshi, S., and S. Shinoda. 2000. Microbial metalloproteases and pathogenesis. Microbes Infect. 2:91-98.
44. Parsek, M. R., and E. P. Greenberg. 2000. Acyl-homoserine lactone quorum sensing in gram-negative bacteria: a signaling mechanism involved in associations with higher organisms. Proc. Natl. Acad. Sci. USA 97:8789-8793.
45. Reik, R., T. Spilker, and J. J. Lipuma. 2005. Distribution of Burkholderia cepacia complex species among isolates recovered from persons with or without cystic fibrosis. J. Clin. Microbiol. 43:2926-2928.
46. Riedel, K., C. Arevalo-Ferro, G. Reil, A. Gorg, F. Lottspeich, and L. Eberl. 2003. Analysis of the quorum-sensing regulon of the opportunistic pathogen Burkholderia cepacia H111 by proteomics. Electrophoresis 24:740-750.
47. Rinderknecht, H., M. C. Geokas, P. Silverman, and B. J. Haverback. 1968. A new ultrasensitive method for the determination of proteolytic activity. Clin. Chim. Acta 21:197-203.
48. Rutherford, K., J. Parkhill, J. Crook, T. Horsnell, P. Rice, M. A. Rajandream, and B. Barrell. 2000. Artemis: sequence visualization and annotation. Bioinformatics 16:944-945.
49. Sajjan, U., C. Ackerley, and J. Forstner. 2002. Interaction of cblA/adhesin-positive Burkholderia cepacia with squamous epithelium. Cell. Microbiol. 4:73-86.
50. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
51. Schagger, H., and G. von Jagow. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379.
52. Sokol, P. A., C. Kooi, R. S. Hodges, P. Cachia, and D. E. Woods. 2000. Immunization with a Pseudomonas aeruginosa elastase peptide reduces severity of experimental lung infections due to P. aeruginosa or Burkholderia cepacia. J. Infect. Dis. 181:1682-1692.
53. Sokol, P. A., D. E. Ohman, and B. H. Iglewski. 1979. A more sensitive plate assay for detection of protease production by Pseudomanas aeruginosa. J. Clin. Microbiol. 9:538-540.
54. Sokol, P. A., U. Sajjan, M. B. Visser, S. Gingues, J. Forstner, and C. Kooi. 2003. The CepIR quorum-sensing system contributes to the virulence of Burkholderia cenocepacia respiratory infections. Microbiology 149:3649-3658.
55. Speert, D. P., D. Henry, P. Vandamme, M. Corey, and E. Mahenthiralingam. 2002. Epidemiology of Burkholderia cepacia complex in patients with cystic fibrosis, Canada. Emerg. Infect. Dis. 8:181-187.
56. Sun, L., R. Z. Jiang, S. Steinbach, A. Holmes, C. Campanelli, J. Forstner, U. Sajjan, Y. Tan, M. Riley, and R. Goldstein. 1995. The emergence of a highly transmissible lineage of cbl+ Pseudomonas (Burkholderia) cepacia causing CF centre epidemics in North America and Britain. Nat. Med. 1:661-666.
57. Tomich, M., C. A. Herfst, J. W. Golden, and C. D. Mohr. 2002. Role of flagella in host cell invasion by Burkholderia cepacia. Infect. Immun. 70:1799-1806.
58. Venturi, V., A. Friscina, I. Bertani, G. Devescovi, and C. Aguilar. 2004. Quorum sensing in the Burkholderia cepacia complex. Res. Microbiol. 155:238-244.
59. Visser, M. B., S. Majumdar, E. Hani, and P. A. Sokol. 2004. Importance of the ornibactin and pyochelin siderophore transport systems in Burkholderia cenocepacia lung infections. Infect. Immun. 72:2850-2857.
60. West, S. E., H. P. Schweizer, C. Dall, A. K. Sample, and L. J. Runyen-Janecky. 1994. Construction of improved Escherichia-Pseudomonas shuttle vectors derived from pUC18/19 and sequence of the region required for their replication in Pseudomonas aeruginosa. Gene 148:81-86.(C. Kooi, B. Subsin, R. Ch)
ABSTRACT
In previous studies we characterized the Burkholderia cenocepacia ZmpA zinc metalloprotease. In this study, we determined that B. cenocepacia has an additional metalloprotease, which we designated ZmpB. The zmpB gene is present in the same species as zmpA and was detected in B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia but was absent from B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina. The zmpB gene was expressed, and ZmpB was purified from Escherichia coli by using the pPROEXHTa His6 Tag expression system. ZmpB has a predicted preproenzyme structure typical of thermolysin-like proteases and is distantly related to Bacillus cereus bacillolysin. ZmpB was expressed as a 63-kDa preproenzyme precursor that was autocatalytically cleaved into mature ZmpB (35 kDa) and a 27-kDa prepropeptide. EDTA, 1,10-phenanthroline, and Zn2+ cations inhibited ZmpB enzyme activity, indicating that it is a metalloprotease. ZmpB had proteolytic activity against -1 proteinase inhibitor, 2-macrogobulin, type IV collagen, fibronectin, lactoferrin, transferrin, and immunoglobulins. B. cenocepacia zmpB and zmpA zmpB mutants had no proteolytic activity against casein and were less virulent in a rat agar bead chronic infection model, indicating that zmpB is involved in B. cenocepacia virulence. Expression of zmpB was regulated by both the CepIR and CciIR quorum-sensing systems.
INTRODUCTION
The Burkholderia cepacia complex is comprised of nine closely related species (8, 9; reviewed in reference 39). B. cepacia complex organisms infect approximately 4 to 7% of cystic fibrosis (CF) patients (12). B. cepacia complex strains may cause a rapid deterioration of lung function and death in some CF patients (27). Strains from all nine species have been isolated from CF patients, but the most prevalent species are B. cenocepacia and B. multivorans, with strains of B. cenocepacia being most commonly isolated from North American CF patients (45, 55). The majority of the transmissible and epidemic B. cepacia complex strains belong to B. cenocepacia (35, 39, 41, 55). Highly transmissible B. cenocepacia strains, such as ET12, Midwest, and PHDC, have been identified in outbreaks in North America and Europe (6, 34). B. cenocepacia ET12 was responsible for the largest B. cepacia complex epidemic affecting CF patients in Canada and the United Kingdom during the late 1980s and early 1990s and has been linked to patient-to-patient transmission (55, 56). B. cenocepacia ET12 strains contain a large number of unique genes (3, 38, 56), although none of these have been directly linked to transmissibility between patients.
Numerous factors that have been implicated in B. cenocepacia virulence have been identified, including protease (11), quorum-sensing systems (1, 30, 54), hemolysin (26), lipopolysaccharide (25), capsule (25), cable pili and 22-kDa adhesin (49), flagella (57), exopolysaccharides (7, 10), siderophores (59), and yfjI, a gene of unknown function (3). We previously identified the zmpA gene and determined that it encodes a zinc metalloprotease that contributes to virulence in chronic lung infections (11). B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia have a zmpA gene and detectable extracellular protease activity, whereas B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina lack zmpA and are protease negative (19). ZmpA has the potential to cause direct tissue damage and to modulate the host immune system, since it has been shown to degrade type IV collagen, fibronectin, -1 proteinase inhibitor, 2-macroglobulin, and gamma interferon (28). ZmpA is expressed as a preproenzyme that is autoproteolytically cleaved into a 36-kDa mature enzyme. It was confirmed to be a zinc metalloprotease, since its activity was inhibited by EDTA and 1,10-phenanthroline (28). Expression of the zmpA gene is regulated by both the CepIR and CciIR quorum-sensing systems (40, 54).
Quorum sensing is a regulatory system that controls expression of target genes in a cell density-dependent manner and in gram-negative bacteria usually involves N-acyl-homoserine lactones (AHLs) as signaling molecules (reviewed in references 44 and 58). AHLs are synthesized by AHL synthases encoded by luxI homologues. The AHLs bind to and activate a response regulator encoded by a luxR homologue that regulates expression of target genes at the level of transcription. Two LuxIR quorum-sensing systems, cepIR and cciIR, have been identified in B. cenocepacia K56-2 (1, 33, 40). CepI and CciI synthesize N-octanoyl-L-homoserine lactone and N-hexanoyl-L-homoserine lactone, although in different ratios (36, 40). The cepIR system is widely distributed throughout the B. cepacia complex (20, 36), whereas the cciIR system is found only in B. cenocepacia ET12 strains that contain the cci genomic island (1).
Protease activity has previously been characterized in B. cenocepacia strains with mutations in zmpA or each of the quorum-sensing genes. B. cenocepacia K56-2 and Pc715j zmpA mutants had significantly less extracellular protease activity than the parent strains; however, the zmpA mutation did not completely eliminate proteolytic activity (11). In fact, the Pc715j zmpA mutant retained approximately 50% of the proteolytic activity of the parent and was equally virulent in the rat agar bead lung infection model (11). B. cenocepacia K56-2 or H111 cepI or cepR mutants produce no detectable protease (24, 32). Interestingly, a K56-2 cciI mutant produced significantly more protease activity than the parent strain, although expression of a zmpA::lacZ fusion was considerably lower in the cciI mutant than the parent strain (40). Taken together, these studies suggested that B. cenocepacia has at least one extracellular protease gene in addition to zmpA and that there are differences between the regulation of zmpA and other protease genes by the cciIR quorum-sensing system. We isolated a spontaneous protease-negative mutant of B. cenocepacia K56-2 that does not express zmpA but has less protease activity than a zmpA mutant, suggesting that it was also deficient in expression of other proteases (data not shown). In the present study, we used this mutant to clone a second metalloprotease gene from B. cenocepacia, which we designated zmpB. We characterized the activity of this protease and demonstrated that it is involved in the virulence of B. cenocepacia and that its expression is regulated by both the CepIR and CciR quorum-sensing systems.
MATERIALS AND METHODS
Strains, plasmids, primers, and growth conditions. Strains and plasmids used in this study are listed in Table 1. Escherichia coli and B. cenocepacia were routinely grown in Luria-Bertani (LB) broth (Invitrogen, Burlington, Ontario, Canada) or on LB agar at 37°C. Bacteria from rat lung homogenates were recovered on Burkholderia cepacia selective agar (21). When appropriate, antibiotics were used at the following concentrations (per milliliter): for B. cenocepacia, 200 μg tetracycline (Tc) and 100 μg trimethoprim (Tp), and for E. coli, 100 μg ampicillin (Ap), 50 μg kanamycin (Km), 15 μg Tc, and 1.5 mg Tp. All chemicals were obtained from Sigma (St. Louis, Mo.) except where indicated.
DNA manipulations. Molecular biology techniques were generally performed as described by Sambrook et al. (50). PCRs were carried out using standard methods as described by the manufacturer, with either Vent DNA polymerase (New England Biolabs, Pickering, Ontario, Canada) for cloning of zmpB or Taq DNA polymerase (Invitrogen) for amplification of a zmpB fragment from B. cepacia complex strains. PCR products or DNA fragments were purified from agarose gels by using a QIAquick gel extraction kit (QIAGEN, Mississauga, Ontario, Canada). DNA sequencing was performed by the University of Calgary Core DNA Sequencing Laboratory. Sequence analysis was performed with DNAMAN software (Lynnon Biosoft, Vaudreuil, Quebec, Canada), and sequences were compared to the unpublished genome sequence of B. cenocepacia J2315 (http://www.sanger.ac.uk/Projects/B_cenocepacia/) by using Artemis software (48).
Cloning of zmpB and construction of zmpB mutants. B. cenocepacia K56-2 genomic DNA was partially digested with Sau3A and cloned into pUCP26 (60) digested with BamHI. The resultant library was electroporated into a spontaneous protease-negative mutant of K56-2 as previously described (14), and transformants were screened for restoration of proteolytic activity on brain heart infusion (BHI)-skim milk agar (53). One positive clone was designated pUCP21D6.
A 2.9-kb PCR fragment containing zmpB was amplified from pUCP21D6 by using primers BS1-21D6F (5'-TGTGCGTCGCGTGCGGCTTC-3') and BS1-21D6R (5'-GTGCGCGACCTGTTCGACGG-3') and cloned into pCR2.1-TOPO (Invitrogen), generating pBS1. For construction of pBS2 and pBS3, antibiotic cassettes (Tpr from p34E-Tp [16] and Tcr from p34S-Tc [15], respectively) were inserted into the pBS1 SalI site after deletion of a 1.4-kb SalI internal fragment.
A 2.1-kb XhoI-HindIII fragment from pBS2 was cloned into the suicide vector pEX18Tc and used to generate K56-2 zmpB by allelic exchange as previously described (22). A 2.9-kb XhoI-HindIII fragment from pBS3 was cloned into pEX18Ap (22) and transformed into B. cenocepacia K56-2-9 (zmpA) (11) to generate K56-2 zmpA zmpB. Allelic exchange mutants were identified by loss of the vector, sucrose resistance, and decreased zones on BHI-skim milk agar. Double-crossover mutants were confirmed by PCR using the primers pBS1-21D6F and BS1-21D6R. For complementation experiments, the 2.9-kb XhoI-HindIII fragment containing zmpB from pBS1 was cloned into pUCP26 (60) to generate pBS6.
Animal studies. The virulence of the K56-2 zmpA, K56-2 zmpB, and K56-2 zmpA zmpB mutants was compared to that of the parent K56-2 by using the agar bead chronic respiratory infection model (4). The number of bacteria and histopathological changes indicating inflammation in the infected lungs were quantified on days 7 and 14 postinfection (p.i.) as previously described (2).
For generation of anti-ZmpB sera, Sprague-Dawley rats (Charles River, Saint Constant, Quebec, Canada) were immunized on days 0 and 21 with recombinant ZmpB (approximately 20 μg) emulsified in monophosphoryl lipid A plus synthetic trehalose dicorynomycolate adjuvant as described by the manufacturer (Corixa Corporation, Hamilton, MT). Rats were bled on day 26 by cardiac puncture and anti-ZmpB antibodies detected by enzyme-linked immunosorbent assay using recombinant ZmpB as the antigen. Production of anti-ZmpA serum was previously described (19).
Expression of the zmpB gene and purification of recombinant ZmpB. The primers ZMPBF (5'-CGGAATTCATGCAGACATCACGCAAAGCATTG) and ZMPBR (5'-CCCAAGCTTCCTGCTGCGGTATTACGACTTCTG) were used to PCR amplify a 1,713-bp fragment containing K56-2 zmpB from the start codon (ATG) to 12 bp beyond the stop codon (TAA). The ZMPBF and ZMPBR primers contain an EcoRI site and a HindIII site (underlined), respectively. The amplified fragment was then digested with EcoRI-HindIII and ligated into pPROEXHTa (Invitrogen) to generate pSCM4.
E. coli DH5(pSCM4) was grown overnight in 10 ml LB supplemented with Ap and used to inoculate 400 ml LB at an initial optical density at 600 nm (OD600) of 0.03. When the cultures reached an OD600 of 0.6, they were induced with 0.7 mM isopropyl--D-thiogalactoside (IPTG) for 3.5 h. Recombinant ZmpB was purified and refolded as previously described for ZmpA (28).
The relative molecular mass and purity of ZmpB were determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (31). The partial N-terminal amino acid sequence of the mature ZmpB was determined from ZmpB separated by SDS-PAGE and electroblotted onto polyvinylidene difluoride (PVDF) membranes (Millipore Corp., Etobicoke, Ontario, Canada). Sequencing was performed by the Protein Microchemistry Laboratory at the University of British Columbia.
To detect ZmpB expression in B. cenocepacia and other B. cepacia complex species, cell-free supernatants were precipitated with trichloroacetic acid to a final concentration of 10% as previously described (11). The proteins were electrophoresed for 30 h at 80 V by Tricine-SDS-14% polyacrylamide gel electrophoresis (51) and transferred to PVDF membranes. ZmpB and ZmpA were detected on the immunoblots with rat antibodies to each protein, using peroxidase-conjugated goat anti-rat immunoglobulin G (IgG) (Kirkegaard & Perry, Gaithersburg, MD).
Protease activity assays. Protease activity was determined on BHI-skim milk agar at 24 and 48 h as previously described (28) or with hide powder azure (47). One unit of protease was defined as the activity that produced a change in the OD595 of 0.1 per hour at 37°C.
The optimal pH for ZmpB proteolytic activity was determined. ZmpB (4 units, 100 ng) was diluted in citrate buffer (25 mM, pH 3 to 6), morpholinoethanesulfonic acid (MES) (25 mM, pH 5 to 6.7), 3-(N-morpholino)propanesulfonic acid (MOPS) (25 mM, pH 6.5 to 8), Tris-HCl (25 mM, pH 8 to 9), or bicarbonate (25 mM, pH 10) and incubated at 37°C for 2 h. The optimum temperature for ZmpB activity was determined in 25 mM MES (pH 5.6). Assays were performed in triplicate with a buffer control at each temperature, using hide powder azure as the substrate.
To identify potential ZmpB substrates, ZmpB (2 units) was incubated for 24 to 48 h at 37°C with -1 proteinase inhibitor (Bayer Corporation, Elkhart, IN), human fibronectin, type IV collagen, lactoferrin, transferrin, IgA, IgG, IgM, rat 2-macroglobulin, and recombinant rat gamma interferon (Sigma) in 25 mM MES, pH 5.6. Recombinant ZmpB and substrate-only controls were included. In addition, substrates were incubated with the proteolytically inactive ZmpA H465G (28) purified in an identical manner as ZmpB to ensure that proteolytic activity was not due to contaminating proteases in the reaction mixtures.
For inhibitor and divalent metal cation studies, ZmpB (4 units, approximately 100 ng) was preincubated with the specified inhibitor(s) and/or divalent metal cation(s) for 30 min at room temperature and transferred to hide powder azure in 25 mM MES (pH 5.6). Neutralization assays with a monoclonal antibody (36-6-8) to ZmpA were performed as previously described (29).
Expression of zmpB::luxCDABE transcriptional fusions. A zmpB::luxCDABE transcriptional fusion was constructed in pMS402 (17). A 0.75-kb DNA fragment containing the zmpB promoter was PCR amplified from K56-2 genomic DNA with BSpzmpBF (5'-ATACTTTCGTATCGGCCAC-3') and BSpzmpBR (5'-ACGCACTTCTACCTGAACCT-3'). The amplified fragment was digested with BamHI-XhoI and ligated into the BamHI-XhoI site upstream of luxCDABE in pMS402, resulting in pBS8. To generate pBS9, a SacI fragment from p34STc (15) containing the tetracycline resistance cassette was ligated into pBS8 digested with XhoI.
Luminescence assays were performed to measure expression of zmpB. Overnight cultures were subcultured in triplicate to an initial OD600 of 0.02 in 20 ml LB and grown at 37°C and 250 rpm. At 16 and 20 h, 150-μl aliquots were transferred into 96-well clear-bottom black plates (9520 Costar; Corning Incorporated, Corning, NY), and the luminescence in counts per second and turbidity at an absorbance of 600 nm were measured using a Wallac Victor2 1420 multilabel counter (Perkin-Elmer Life Sciences, Boston, Mass.). Expression levels of zmpB were determined by relative luminescence.
Distribution of the zmpB gene in the B. cepacia complex. To determine the distribution of the zmpB gene in Burkholderia species, primers ZPBDTF (GCAGACATCACGCAAAGCATTG) and ZPBDTR (GCAGCTCTTGTTGTACGCGTTG) were used to PCR amplify a 1.7-kb zmpB fragment. The amplification conditions were 95°C for 7 min; followed by 30 cycles of 95°C for 1 min, 56°C for 30 s, and 72°C for 2 min; followed by 72°C for 20 min.
Southern hybridization analysis was also performed on genomic DNA isolated from representative strains digested with NotI. A 0.8-kb MluI fragment encoding from Arg-35 to Asn-307 of the 323-amino-acid mature ZmpB was labeled with 32P by random priming as described by the manufacturer (GE Healthcare) and used as a probe. Blots were hybridized with the probe at 65°C and washed at 72°C.
RESULTS
Identification of the zmpB gene. The plasmid pUCP21D6 with a 6,780-bp insert containing a putative metalloprotease gene was identified by complementation of a spontaneous protease-negative mutant that does not express zmpA (data not shown) with a B. cenocepacia K56-2 genomic library. Analysis of this insert by using the unpublished annotation of B. cenocepacia J2315 at the Sanger Institute (http://www.sanger.ac.uk/Projects/B_cenocepacia/) and Artemis software (48) revealed five open reading frames encoding two transcriptional regulators of the AraC family (BCAM2305 and BCAM2306), a thermolysin metalloprotease (BCAM2307), a leucyl aminopeptidase (BCAM2308), and a hypothetical protein (BCAM2309). BCAM2307, encoding a putative thermolysin-like metalloprotease belonging to the M4 family of peptidases, was designated zmpB, and its corresponding protein was designated ZmpB. Alignment analysis of ZmpB with the proteases most related to it, bacillolysin from Bacillus cereus ATCC 14579 (29% identity) and thermolysin peptidase M4 from Bacillus anthracis A2012 (27% identity), indicated that the HExxH active-site motif and amino acids downstream of the active site (including GxxxExxxD, where E is a putative zinc ligand) are conserved. Functional domain analysis using the Conserved Domain Database (http://www.ncbi.nlm.nih.gov/Structure/cdd) demonstrated that ZmpB is also functionally related to Pseudomonas aeruginosa LasB and has a prepropeptide structure typical of secreted metalloproteases. The precursor to ZmpB has a theoretical pI of 5.84 and a molecular mass of 60547 Da (http://us.expasy.org.). The pre sequence has a hydrophobic transmembrane domain at amino acids 7 to 26 (http://www.sbc.su.se/miklos/DAS) and a predicted SignalP recognition cleavage site 24TVA. The N-terminal amino acid sequence (FVFRPDPLS) was determined with a recombinant ZmpB peptide described below. Based on this amino acid sequence, the mature ZmpB has a predicted molecular mass of 34.9 kDa and pI of 5.08.
Characterization of K56-2 zmpB and K56-2 zmpA zmpB mutants. Mutation of the zmpB gene in K56-2 and K56-2-9 zmpA was performed by insertional inactivation of zmpB with a Tpr or Tcr cassette, resulting in K56-2 zmpB and K56-2 zmpA zmpB, respectively. The extracellular protein profiles were analyzed by Tricine-SDS-14% PAGE (Fig. 1). K56-2 and K56-2-9 zmpA expressed a 35-kDa protein, which corresponded to the predicted molecular mass of ZmpB. The 35-kDa protein was not detected in K56-2 zmpB and K56-2 zmpA zmpB supernatants (Fig. 1A). The 35-kDa proteins of K56-2, K56-2-9 zmpA, and K56-2 zmpB(pBS6) reacted with anti-rat ZmpB antibody (Fig. 1B), indicating that pBS6 complemented K56-2 zmpB. The 36-kDa peptides present in K56-2, K56-2 zmpB, and K56-2 zmpB (pBS6) correspond to the molecular mass of ZmpA (29, 42) and reacted with anti-ZmpA (Fig. 1C). Anti-ZmpA and anti-ZmpB antibodies did not cross-react with the heterologous proteases, suggesting that these two proteases are antigenically distinct.
FIG. 1. Expression of ZmpA and ZmpB in K56-2 zmpB and K56-2 zmpA zmpB mutants. Supernatant proteins were precipitated with 10% trichloroacetic acid, separated by Tricine-14% SDS-PAGE, and blotted to PVDF membranes. A. Coomassie blue-stained gel. B. Western blot reacted with anti-ZmpB. C. Western blot reacted with anti-ZmpA. Lanes 1, K56-2; lanes 2, K56-2 zmpB; lanes 3, K56-2 zmpB(pBS6); lanes 4, K56-2 zmpA zmpB; lanes 5, K56-2 zmpA; lanes 6, recombinant ZmpB; lanes 7, recombinant ZmpA. Solid arrow, ZmpA; open arrow, ZmpB.
The proteolytic activities of K56-2, K56-2 zmpB, and K56-2 zmpA zmpB mutants were determined on BHI-skim milk agar (Table 2). K56-2 had significantly more protease activity than K56-2 zmpB and K56-2 zmpA zmpB. K56-2 zmpB(pBS6) had zones comparable in size to those of K56-2, indicating that pBS6, containing zmpB, complemented the protease-negative phenotype of K56-2 zmpB. The protease activity of K56-2-9 zmpA was similar to that previously reported (11). These results indicate that zmpB is responsible for the majority of the protease activity of B. cenocepacia K56-2. Complementation of K56-2 zmpB with pBS6 did not result in increased protease activity above that of the wild type despite the potential for increased amounts of ZmpB protein due to the multicopy plasmid. Similar results were reported for complementation of zmpA mutants (11).
The effect of a zmpB mutation on virulence was examined using a chronic respiratory infection model. Previously, we reported that the K56-2-9 zmpA mutant was less persistent in lungs and manifested less histopathology than the parent K56-2 (11). Similar results were obtained in this study in that the number of K56-2-9 zmpA or K56-2 zmpA zmpB organisms recovered from the lungs at day 14 p.i. was approximately three log units lower than the number of K56-2 organisms recovered (Table 3). Three of the four rats infected with K56-2-9 zmpA were completely cleared of bacteria by day 14 p.i. Bacteria recovered from the fourth rat were determined by sequence analysis to be revertants. The numbers of K56-2 zmpB organisms recovered from the lungs were similar to those of K56-2 at both 7 and 14 days p.i. (Table 3), suggesting that zmpA plays a role in bacterial persistence but that zmpB does not.
Both zmpA and zmpB contribute to lung pathology. The lungs from the rats infected with the K56-2 zmpA, K56-2 zmpB, or K56-2 zmpA zmpB protease mutant had significantly fewer histopathological changes than the lungs from rats infected with the K56-2 parent (Table 3), indicating that both proteases contribute to the maximal virulence of B. cenocepacia K56-2 in this infection model. There was no significant difference in the quantitative histopathological changes in the K56-2 zmpA zmpB double mutant and the zmpA or zmpB single mutant.
Purification and characteristics of recombinant ZmpB. The zmpB gene was cloned into pPROEXHTa to construct pSCM4 and was expressed as an N-terminal His-tagged protein. E. coli DH5(pSCM4) was proteolytically active on BHI-skim milk agar, with zones of approximately 2 to 4 mm after 48 h, in contrast to E. coli DH5(pPROEXHTa), which displayed no proteolytic activity. The zmpB gene was expressed as a 63-kDa protein in the cell lysate after IPTG induction for 3.5 h (Fig. 2, lane 1). The mass of 63 kDa corresponded to the predicted molecular mass of the His6-tagged ZmpB (63.4 kDa). When the cell lysate was fractionated and analyzed by SDS-PAGE, the 63-kDa protein was recovered almost exclusively in the precipitate as insoluble inclusion bodies (Fig. 2, lane 3). When the inclusion bodies were solubilized in 6 M guanidine-HCl and applied to an Ni-nitrilotriacetic acid (NTA) affinity column to purify and refold the 63-kDa protein, two new peptides (35 kDa and 28 kDa) were identified in the eluant (Fig. 2, lane 4). The N-terminal sequence of the 35-kDa peptide was determined to be FVFRPDPLS, which is present in the C-terminal portion of the 63-kDa His6-tagged ZmpB, and this mass corresponds to the predicted mass of 34.9 kDa that would be obtained for a peptide with this N-terminal sequence. Therefore, the 35-kDa peptide contains the active site and is the mature protease. The 28-kDa peptide has the predicted mass for the prepropeptide and the His tag.
FIG. 2. Expression and purification of ZmpB. Lane M, molecular weight markers (in thousands); lane 1, cell lysate after induction with IPTG; lane 2, uninduced crude cell extract; lane 3, purified inclusion bodies; lane 4, purified recombinant ZmpB after refolding and elution from an Ni-NTA column. Solid arrow, His-prepro-ZmpB fusion protein; open arrow, indicates the His-prepropeptide. The mature ZmpB is indicated.
ZmpB eluted from the Ni-NTA column was proteolytically active against hide powder azure, with 50 ng containing approximately 5 U of activity. The effect of protease inhibitors or divalent cations on ZmpB protease activity was examined. ZmpB activity was inhibited by EDTA and 1,10-phenanthroline, which chelate Zn2+, but not by phenylmethylsulfonyl fluoride or phosphoramidon (Table 4). ZmpB was not inhibited by 1,7-phenanthroline, an isomer of 1,10-phenanthroline that does not bind Zn2+. In addition, 1 mM Zn2+ abrogated ZmpB activity, but 1 mM CaCl2 had no effect (Table 4). Co2+ and Ni2+ also significantly reduced ZmpB activity. Monoclonal antibody 36-6-8, which previously was shown to inhibit ZmpA, did not inhibit ZmpB activity (29). The A595 when ZmpB was preincubated with monoclonal antibody 36-6-8 was 0.7 ± 0.2, compared with 0.7 ± 0.1 in control ascites fluid.
The optimal pH range for ZmpB activity was between pH 5.6 and 7.2, depending on the buffers employed (data not shown). ZmpB was active over a broad pH range, although activity was dramatically decreased at pH values below 4.5 or above 8.0. ZmpB was active between 28 and 60°C, with an optimum temperature of 50°C (data not shown). Activity was significantly decreased at temperatures above 70°C.
ZmpB activity against host proteins. The ability of ZmpB to degrade a number of biologically important substrates which might account for its role in virulence was determined by incubating ZmpB with the substrate and determining proteolysis by SDS-PAGE (Fig. 3). ZmpB was able to digest the human plasma protease inhibitors -1 proteinase inhibitor and 2-macroglobulin. ZmpB also digested type IV collagen, fibronectin, transferrin, lactoferrin, and immunoglobulins. Collagen, fibronectin, -1 proteinase inhibitor, and 2-macroglobulin were cleaved into numerous peptides, indicating that ZmpB has broad specificity. Transferrin, lactoferrin, and the immunoglobulins IgA, IgG, and IgM were also cleaved into several peptides but were less extensively degraded by ZmpB than collagen, fibronectin, -1 proteinase inhibitor, and 2-macroglobulin.
FIG. 3. Substrate specificity of ZmpB. Substrates were digested with ZmpB, ZmpAH465G (which has no proteolytic activity), or in the absence of enzyme (no protease) for 24 to 48 h. Proteins were separated by SDS-PAGE and stained with Coomassie blue. Collagen, fibronectin, and 2-macroglobulin were electrophoresed by SDS-7.5% PAGE. All other substrates were electrophoresed by SDS-12.5% PAGE.
In order to ensure that the ZmpB activity was not due to the presence of a contaminating E. coli protease, these substrates were digested with a recombinant inactive form of ZmpA, ZmpA H465G (28), that was purified using the same protocol. ZmpA H465G had no activity against any of the substrates, confirming that the cleavage of all substrates was not due to the presence of contaminating proteases in the reaction mixtures or proteases that might copurify from E. coli.
Quorum-sensing regulation of zmpB expression. Previously, zmpA had been shown to be regulated by the cepIR and cciIR quorum-sensing systems (40, 54). To determine if zmpB expression was regulated by cepIR or cciIR, the zmpB promoter region was cloned upstream of the luxCDABE operon in pMS402 (17). The resulting plasmid, pBS9, was transformed into K56-2, K56-dI2 cepI, K56-R2 cepR, K56-2 cciI, K56-2 cciR, K56 cciIR, and K56-2 cepR cciIR. Expression of zmpB::luxCDABE was reduced to almost background levels in K56-dI2(pBS9) and K56-R2(pBS9) and was significantly reduced in K56-2 cciR(pBS9) and K56-2 cepR cciIR(pBS9) (Fig. 4). Interestingly, zmpB::luxCDABE was significantly increased in K56-2 cciI(pBS9). Expression in K56-2 cciIR(pBS9) was similar to that in the parent. These data suggest that zmpB expression is regulated by both cepIR and cciIR.
FIG. 4. Regulation of zmpB by the cepIR and cciIR quorum-sensing genes. Plasmid pBS9 containing a zmpB::luxCDABE fusion was introduced into K56-2 quorum-sensing mutants. Cultures were grown for 20 h, and luminescence in counts per second (CPS) and turbidity at an absorbance of 600 nm were measured using a Wallac Victor2 1420 multilabel counter. Values shown are the mean relative CPS (±standard deviation) for triplicate cultures and are representative of three independent experiments. , significantly different from result for K56-2 (P < 0.05 by analysis of variance).
Distribution of the zmpB gene in Burkholderia spp. To investigate the distribution of the zmpB gene in Burkholderia species, primers ZPBDTF and ZPBDTR were used to PCR amplify the 1.7-kb zmpB gene in 43 strains representing the B. cepacia complex. A 1.7-kb amplicon was obtained from all strains of B. cepacia, B. cenocepacia, B. ambifaria, and B. pyrrocinia examined but not from strains of B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina (data not shown). Southern blot hybridization analysis was also performed to determine which species contained a zmpB gene (Fig. 5A). B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia had a 0.8-kb NotI band that hybridized to the zmpB probe as strongly as that of the positive control strain K56-2 (Fig. 5A). The probe did not hybridize to B. multivorans, B. vietnamiensis, B. dolosa, or B. anthina, suggesting that these species do not contain a gene related to zmpB, although it is possible that these species could contain a related gene with less homology than would be detected by Southern hybridization.
FIG. 5. Distribution of ZmpB in the B. cepacia complex. A. Southern hybridization analysis of the zmpB gene. A 0.8-kb 32P-labeled MluI fragment encoding from Arg-35 to Asn-307 of the 323-amino-acid mature ZmpB from K56-2 was used to probe NotI-digested genomic DNA. Lane 1, K56-2; lane 2, B. cepacia LMG17997; lane 3, B. multivorans LMG13010; lane 4, B. cenocepacia J2315; lane 5, B. stabilis LMG14294; lane 6, B. stabilis C7322; lane 7, B. vietnamiensis PC259; lane 8, B. dolosa LMG18943; lane 9, B. ambifaria Cep996; lane 10, B. anthina LMG20980; lane 11, B. pyrrocinia LMG21822. B. Detection of ZmpB by Western blotting of culture supernatants. Trichloroacetic acid-precipitated culture supernatants were separated by Tricine 14%-PAGE and blotted onto PVDF membrane. Blots were reacted with anti-ZmpB. Lane 1, K56-2; lane 2, B. cepacia LMG17997; lane 3, B. multivorans LMG13010; lane 4, B. cenocepacia C6433; lane 5, B. stabilis LMG18888; lane 6, B. vietnamiensis Pc259; lane 7, B. dolosa LMG18943; lane 8, B. ambifaria CEP996; lane 9, B. anthina LMG20980; lane 10, B. pyrrocinia LMG 21822; lane 11, recombinant ZmpB.
To determine if the presence of zmpB correlated with extracellular protease activity, we examined the activity of 43 strains on BHI-skim milk agar. The majority (18 out of 23) of the zmpB-positive strains had extracellular protease activity, while none of the zmpB-negative strains produced detectable extracellular protease activity. These data suggest that the strains in which zmpB was not detectable by PCR or Southern hybridization do not contain a related gene. ZmpB was easily detected only in B. cenocepacia concentrated culture supernatants by Western blot analysis (Fig. 5B). Possibly ZmpB was not expressed and/or secreted in the other zmpB-positive B. cepacia complex strains under the conditions employed, or ZmpB was secreted but the anti-ZmpB polyclonal antibody employed did not react with the ZmpB from these strains.
DISCUSSION
Metalloproteases are widely distributed in nature and belong to four superfamilies or clans. All bacterial zinc metalloproteases belong to the zincins superfamily, which contains a conserved HEXXH motif. The two histidines are zinc ligands, and the glutamic acid is involved in enzymatic activity. The zincins can be subdivided, based on the position of the third zinc ligand (43), into at least 10 families, with 3 of the families containing bacterial metalloproteases: thermolysin, serralysin, and neurotoxins.
B. cenocepacia ZmpB is a novel thermolysin-like protease that has only 28% identity with its closest homologue, bacillolysin from Bacillus cereus, and shares only 10.3% identity with B. cenocepacia ZmpA. ZmpB has a preproenzyme structure that is typical of the M4 family (http://merops.sanger.ac.uk/) of metalloproteases, with conservation of functional regions such as the HExxH active-site motif as well as other conserved amino acids downstream of the active site, including GxxxExxxD. De Kreij et al. (13) found that there was a great deal of sequence variability among various thermolysin-like proteases but that as long as a small subset of amino acids was conserved (e.g., active site residues and zinc ligands), the group was extremely homogeneous in terms of catalysis. B. cenocepacia ZmpB is inhibited by EDTA and 1,10-phenanthroline, indicating that it is a metalloprotease. ZmpB is thermostable and active over a broad neutral pH range, similar to other bacterial thermolysin-like metalloproteases. ZmpB activity was relatively unaffected by Ca2+ and Mg2+ but completely abrogated by excess Zn2+, as has been reported with thermolysin (23). Co2+ and to lesser degree Mn2+ could substitute for Zn2+ in thermolysin (23), which is also suggested by our findings for ZmpB. Ni2+ has not been shown to inhibit thermolysin, but Ni2+, Zn2+, and Cu2+ have been shown to inhibit a Vibrio vulnificus broad-specificity metalloprotease (5). Interestingly, ZmpB was not inhibited by phosphoramidon, a peptide inhibitor of metalloproteases that is related to B. thermoproteolyticus thermolysin. B. cenocepacia ZmpA is also unaffected by phosphoramidon (28). Phosphoramidon is a peptide that binds to a specific sequence in the thermolysin active-site region. Sequence differences in this region have been correlated with reduced phosphoramidon sensitivity (13).
ZmpB was shown to be a broad-spectrum protease capable of degrading substrates involved in tissue integrity and host defense. Most proteases are inhibited by 2-macroglobulin, but both ZmpB and ZmpA (28) are able to digest 2-macroglobulin and -1 proteinase inhibitor, indicating that B. cenocepacia produces two extracellular proteases that can inactivate the major mammalian protease inhibitors. Inactivation of -1 proteinase inhibitor in vivo may disrupt the -1 proteinase/-1 proteinase inhibitor balance and lead to increased tissue damage and inflammation. Inactivation of 2-macroglobulin could result in increased B. cenocepacia dissemination and lead to septicemia. Both ZmpB and ZmpA can degrade host tissue components, including collagen and fibronectin. Although ZmpB and ZmpA have activity against many of the same substrates, each enzyme has a different specificity. ZmpA cleaved -1 proteinase inhibitor specifically at D36 (D36TSHHDQD) (28), whereas ZmpB cleaved -1 proteinase inhibitor into a number of fragments (Fig. 3). ZmpA specifically cleaved 2-macroglobulin into major polypeptides of 95 and 86 kDa (28). ZmpB cleaved 2-macroglobulin into numerous peptides (Fig. 3).
ZmpB was shown to cleave immunoglobulins (IgA, IgG, and IgM), transferrin, and lactoferrin, which could also increase the ability of B. cenocepacia to evade host defenses. Although we previously cited unpublished data (52) that suggested that ZmpA had activity against these substrates, recombinant and native ZmpA were subsequently shown not to degrade these substrates (28). It is likely that the protease preparations used in the unpublished study contained a mixture of both ZmpA and ZmpB proteases. ZmpB digested lactoferrin, transferrin, and immunoglobulins into markedly fewer peptides than 2-macroglobulin, collagen, and fibronectin, suggesting that there is some cleavage site specificity. ZmpA was shown to digest gamma interferon (28). Because ZmpB had significant proteolytic activity against the albumin present in large amounts in the gamma interferon preparation used, it was not possible to conclusively distinguish between gamma interferon and albumin cleavage products in this assay.
ZmpA was previously shown to be necessary for B. cenocepacia K56-2 to cause persistent respiratory infections in rats (11). Similar results were obtained in the present study, in that the numbers of K56-2 zmpA and K56-2 zmpA zmpB organisms recovered from infected lungs were approximately 3 log units lower than in the parent strain. In contrast, K56-2 zmpB was able to survive at parental levels in the rat lung, indicating that ZmpB is not required for persistence in this strain. Infection with any of the mutants resulted in significantly less lung inflammation than infection with the parent strain. There was no significant difference between the quantitative histopathology changes in lungs infected with the mutants. The decreased inflammation observed in lungs infected with zmpA mutants compared to the parent is at least partially due to the decreased numbers of bacteria present in the lung, whereas the decreased inflammation observed with the zmpB mutant is likely due to the absence of this enzyme. Although a zmpA mutation in K56-2 resulted in decreased virulence, a Pc715j zmpA mutant was virulent in the lung infection model (11). Unfortunately, we were not able to construct a zmpB mutation in either Pc715j or Pc715j zmpA to determine its effects on virulence.
Although zmpA and zmpB are not genetically linked and are in fact located on different chromosomes, they are present in the same B. cepacia complex species. Both genes are present in B. cepacia, B. cenocepacia, B. stabilis, B. ambifaria, and B. pyrrocinia but are absent from B. multivorans, B. vietnamiensis, B. dolosa, and B. anthina (19). Presence or absence of these genes is conserved within each species. Highly (>95%) conserved ZmpB homologues are present in the genomes of B. cepacia R18194, B. cenocepacia J2315, B. cenocepacia HI2424, and B. ambifaria AMMD.
Some B. cepacia complex strains were previously reported to have no detectable protease activity despite having the zmpA gene (19). Others had similar levels of protease activity, but the induction of protease production appeared to be much slower. Interestingly, all of these zmpA-positive, protease-deficient strains are also zmpB positive, suggesting that zmpB may also be poorly expressed or nonfunctional in these strains. These data suggest that zmpA and zmpB share similar regulatory mechanisms. It is also possible that these enzymes act synergistically, since they have different cleavage properties. The zmpB mutant has no activity against casein in the BHI-skim milk agar assay, and the zmpA mutant has approximately 20% of the activity of the parent, suggesting that these enzymes may work together to degrade casein, resulting in zones of clearance. The histopathologies observed in lungs of infected rats were similar for animals infected with either the zmpA, zmpB, or zmpA zmpB double mutant, again suggesting that the activities of these enzymes are not additive, since knocking out either gene results in a similar reduction in virulence.
Previously, we have reported that mutations in cepI, cepR, cciI, or cciR result in decreased zmpA expression in K56-2 (40, 54). In this study, we determined that zmpB expression was also markedly reduced in cepI, cepR, cciR, and cepR cciIR mutants. The expression of the zmpB::luxCDABE fusion was not above the background fluorescence obtained with the pMS402 vector control in the cepI or cepR mutants, indicating that the cepIR quorum-sensing genes are required for expression of both zmpB and zmpA. These results correlate with the observed absence of protease activity in cepI and cepR mutants (32, 40). The cciR and cepR cciR mutants were also previously shown to have decreased protease activity compared to the parent strain, which correlates with the zmpB expression data obtained in this study. Interestingly, the cciI mutant expressed zmpB::luxCDABE at much higher levels than the parent or any of the other mutants. The increase in zmpB expression corresponds to the increased protease activity previously observed in the cciI mutant on skim milk agar (40). The zones of protease activity in the cciI mutant could be restored to parental levels by the addition of 2.5 nM N-hexanoyl-L-homoserine lactone, which is the major AHL synthesized by cciI. These data suggest that the protease activity observed in the cciI mutant is due to the increased expression of zmpB and that the observed decrease in zmpA expression has little impact on total protease activity. It is interesting that the presence of zmpB on a multicopy plasmid does not result in increased protease zone sizes similar to that observed in the cciI mutant but only restores protease activity to the parental level. This finding suggests that zmpB copy number is not sufficient for increased protease expression or activity but that other factors, such as the balance of AHL signals, have a significant influence on the amount of protease produced. Mutations in cciI and cciR result in different effects on zmpB expression and protease activity. These mutants also have different swarming phenotypes, suggesting that other genes must also be differentially expressed in cciI and cciR mutants (40).
The ZmpB precursor is processed into mature ZmpB and a propeptide with a predicted molecular mass of 23.1 kDa and a pI of 8.8. The propeptide mass and pI correspond to a peptide identified in a proteome comparison between B. cenocepacia H111 and an H111 cepI mutant (46). The QS8 ID spot was identified as an intracellular thermolysin metallopeptidase and presumed to be a degradation product of a precursor metalloprotease. The N-terminal sequence of QS8 was determined to be DTKVSN, which corresponds to the predicted N terminus of the ZmpB propeptide after removal of the pre-signal peptide (46). Peptide QS8 was found to be approximately 20-fold up-regulated in the H111 parent compared to the cepI mutant, which is consistent with our data indicating that zmpB is positively regulated by the cepIR system.
In summary, we have shown that B. cenocepacia has two zinc metalloproteases that contribute to virulence and have the potential to interfere with host defense mechanisms. Further studies are under way in our laboratory to determine the in vivo activities of these proteases and their mechanisms of regulation.
ACKNOWLEDGMENTS
This work was supported by grants from the Canadian Institutes of Health Research and the Canadian Cystic Fibrosis Foundation to P.A.S.
We thank members of the Sokol laboratory for helpful discussions and critical review of the manuscript.
FOOTNOTES
These authors contributed equally to this study.
REFERENCES
1. Baldwin, A., P. A. Sokol, J. Parkhill, and E. Mahenthiralingam. 2004. The Burkholderia cepacia epidemic strain marker is part of a novel genomic island encoding both virulence and metabolism-associated genes in Burkholderia cenocepacia. Infect. Immun. 72:1537-1547.
2. Bernier, S. P., L. Silo-Suh, D. E. Woods, D. E. Ohman, and P. A. Sokol. 2003. Comparative analysis of plant and animal models for characterization of Burkholderia cepacia virulence. Infect. Immun. 71:5306-5313.
3. Bernier, S. P., and P. A. Sokol. 2005. Use of suppression-subtractive hybridization to identify genes in the Burkholderia cepacia complex that are unique to Burkholderia cenocepacia. J. Bacteriol. 187:5278-5291.
4. Cash, H. A., D. E. Woods, B. McCullough, W. G. Johanson, Jr., and J. A. Bass. 1979. A rat model of chronic respiratory infection with Pseudomonas aeruginosa. Am. Rev. Respir. Dis. 119:453-459.
5. Chang, A. K., H. Y. Kim, J. E. Park, P. Acharya, I. S. Park, S. M. Yoon, H. J. You, K. S. Hahm, J. K. Park, and J. S. Lee. 2005. Vibrio vulnificus secretes a broad-specificity metalloprotease capable of interfering with blood homeostasis through prothrombin activation and fibrinolysis. J. Bacteriol. 187:6909-6916.
6. Chen, J. S., K. A. Witzmann, T. Spilker, R. J. Fink, and J. J. LiPuma. 2001. Endemicity and inter-city spread of Burkholderia cepacia genomovar III in cystic fibrosis. J. Pediatr. 139:643-649.
7. Chung, J. W., E. Altman, T. J. Beveridge, and D. P. Speert. 2003. Colonial morphology of Burkholderia cepacia complex genomovar III: implications in exopolysaccharide production, pilus expression, and persistence in the mouse. Infect. Immun. 71:904-909.
8. Coenye, T., P. Vandamme, J. R. Govan, and J. J. LiPuma. 2001. Taxonomy and identification of the Burkholderia cepacia complex. J. Clin. Microbiol. 39:3427-3436.
9. Coenye, T., P. Vandamme, J. J. LiPuma, J. R. Govan, and E. Mahenthiralingam. 2003. Updated version of the Burkholderia cepacia complex experimental strain panel. J. Clin. Microbiol. 41:2797-2798.
10. Conway, B. A., K. K. Chu, J. Bylund, E. Altman, and D. P. Speert. 2004. Production of exopolysaccharide by Burkholderia cenocepacia results in altered cell-surface interactions and altered bacterial clearance in mice. J. Infect. Dis. 190:957-966.
11. Corbett, C. R., M. N. Burtnick, C. Kooi, D. E. Woods, and P. A. Sokol. 2003. An extracellular zinc metalloprotease gene of Burkholderia cepacia. Microbiology 149:2263-2271.
12. Cystic Fibrosis Foundation. 2002. Cystic Fibrosis Foundation patient registry annual data report. Cystic Fibrosis Foundation, Bethesda, Md.
13. de Kreij, A., G. Venema, and B. van den Burg. 2000. Substrate specificity in the highly heterogeneous M4 peptidase family is determined by a small subset of amino acids. J. Biol. Chem. 275:31115-31120.
14. Dennis, J. J., and P. A. Sokol. 1995. Electrotransformation of Pseudomonas. Methods Mol. Biol. 47:125-133.
15. Dennis, J. J., and G. J. Zylstra. 1998. Plasposons: modular self-cloning minitransposon derivatives for rapid genetic analysis of gram-negative bacterial genomes. Appl. Environ. Microbiol. 64:2710-2715.
16. DeShazer, D., and D. E. Woods. 1996. Broad-host-range cloning and cassette vectors based on the R388 trimethoprim resistance gene. BioTechniques 20:762-764.
17. Duan, K., C. Dammel, J. Stein, H. Rabin, and M. G. Surette. 2003. Modulation of Pseudomonas aeruginosa gene expression by host microflora through interspecies communication. Mol. Microbiol. 50:1477-1491.
18. Figurski, D. H., and D. R. Helinski. 1979. Replication of an origin-containing derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl. Acad. Sci. USA 76:1648-1652.
19. Gingues, S., C. Kooi, M. B. Visser, B. Subsin, and P. A. Sokol. 2005. Distribution and expression of the ZmpA metalloprotease in the Burkholderia cepacia complex. J. Bacteriol. 187:8247-8255.
20. Gotschlich, A., B. Huber, O. Geisenberger, A. Togl, A. Steidle, K. Riedel, P. Hill, B. Tummler, P. Vandamme, B. Middleton, M. Camara, P. Williams, A. Hardman, and L. Eberl. 2001. Synthesis of multiple N-acylhomoserine lactones is wide-spread among the members of the Burkholderia cepacia complex. Syst. Appl. Microbiol. 24:1-14.
21. Henry, D., M. Campbell, C. McGimpsey, A. Clarke, L. Louden, J. L. Burns, M. H. Roe, P. Vandamme, and D. Speert. 1999. Comparison of isolation media for recovery of Burkholderia cepacia complex from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 37:1004-1007.
22. Hoang, T. T., R. R. Karkhoff-Schweizer, A. J. Kutchma, and H. P. Schweizer. 1998. A broad-host-range Flp-FRT recombination system for site-specific excision of chromosomally-located DNA sequences: application for isolation of unmarked Pseudomonas aeruginosa mutants. Gene 212:77-86.
23. Holland, D. R., A. C. Hausrath, D. Juers, and B. W. Matthews. 1995. Structural analysis of zinc substitutions in the active site of thermolysin. Protein Sci. 4:1955-1965.
24. Huber, B., K. Riedel, M. Hentzer, A. Heydorn, A. Gotschlich, M. Givskov, S. Molin, and L. Eberl. 2001. The cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility. Microbiology 147:2517-2528.
25. Hunt, T. A., C. Kooi, P. A. Sokol, and M. A. Valvano. 2004. Identification of Burkholderia cenocepacia genes required for bacterial survival in vivo. Infect. Immun. 72:4010-4022.
26. Hutchison, M. L., I. R. Poxton, and J. R. Govan. 1998. Burkholderia cepacia produces a hemolysin that is capable of inducing apoptosis and degranulation of mammalian phagocytes. Infect. Immun. 66:2033-2039.
27. Isles, A., I. Maclusky, M. Corey, R. Gold, C. Prober, P. Fleming, and H. Levison. 1984. Pseudomonas cepacia infection in cystic fibrosis: an emerging problem. J. Pediatr. 104:206-210.
28. Kooi, C., C. R. Corbett, and P. A. Sokol. 2005. Functional analysis of the Burkholderia cenocepacia ZmpA metalloprotease. J. Bacteriol. 187:4421-4429.
29. Kooi, C., A. Cox, P. Darling, and P. A. Sokol. 1994. Neutralizing monoclonal antibodies to an extracellular Pseudomonas cepacia protease. Infect. Immun. 62:2811-2817.
30. Kothe, M., M. Antl, B. Huber, K. Stoecker, D. Ebrecht, I. Steinmetz, and L. Eberl. 2003. Killing of Caenorhabditis elegans by Burkholderia cepacia is controlled by the cep quorum-sensing system. Cell. Microbiol. 5:343-351.
31. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.
32. Lewenza, S., B. Conway, E. P. Greenberg, and P. A. Sokol. 1999. Quorum sensing in Burkholderia cepacia: identification of the LuxRI homologs CepRI. J. Bacteriol. 181:748-756.
33. Lewenza, S., and P. A. Sokol. 2001. Regulation of ornibactin biosynthesis and N-acyl-L-homoserine lactone production by CepR in Burkholderia cepacia. J. Bacteriol. 183:2212-2218.
34. Lipuma, J. J. 2005. Update on the Burkholderia cepacia complex. Curr. Opin. Pulm. Med. 11:528-533.
35. LiPuma, J. J., T. Spilker, L. H. Gill, P. W. Campbell III, L. Liu, and E. Mahenthiralingam. 2001. Disproportionate distribution of Burkholderia cepacia complex species and transmissibility markers in cystic fibrosis. Am. J. Respir. Crit. Care Med. 164:92-96.
36. Lutter, E., S. Lewenza, J. J. Dennis, M. B. Visser, and P. A. Sokol. 2001. Distribution of quorum-sensing genes in the Burkholderia cepacia complex. Infect. Immun. 69:4661-4666.
37. Mahenthiralingam, E., T. Coenye, J. W. Chung, D. P. Speert, J. R. Govan, P. Taylor, and P. Vandamme. 2000. Diagnostically and experimentally useful panel of strains from the Burkholderia cepacia complex. J. Clin. Microbiol. 38:910-913.
38. Mahenthiralingam, E., D. A. Simpson, and D. P. Speert. 1997. Identification and characterization of a novel DNA marker associated with epidemic Burkholderia cepacia strains recovered from patients with cystic fibrosis. J. Clin. Microbiol. 35:808-816.
39. Mahenthiralingam, E., T. A. Urban, and J. B. Goldberg. 2005. The multifarious, multireplicon Burkholderia cepacia complex. Nat. Rev. Microbiol. 3:144-156.
40. Malott, R. J., A. Baldwin, E. Mahenthiralingam, and P. A. Sokol. 2005. Characterization of the cciIR quorum-sensing system in Burkholderia cenocepacia. Infect. Immun. 73:4982-4992.
41. McDowell, A., E. Mahenthiralingam, J. E. Moore, K. E. Dunbar, A. K. Webb, M. E. Dodd, S. L. Martin, B. C. Millar, C. J. Scott, M. Crowe, and J. S. Elborn. 2001. PCR-based detection and identification of Burkholderia cepacia complex pathogens in sputum from cystic fibrosis patients. J. Clin. Microbiol. 39:4247-4255.
42. McKevitt, A. I., S. Bajaksouzian, J. D. Klinger, and D. E. Woods. 1989. Purification and characterization of an extracellular protease from Pseudomonas cepacia. Infect. Immun. 57:771-778.
43. Miyoshi, S., and S. Shinoda. 2000. Microbial metalloproteases and pathogenesis. Microbes Infect. 2:91-98.
44. Parsek, M. R., and E. P. Greenberg. 2000. Acyl-homoserine lactone quorum sensing in gram-negative bacteria: a signaling mechanism involved in associations with higher organisms. Proc. Natl. Acad. Sci. USA 97:8789-8793.
45. Reik, R., T. Spilker, and J. J. Lipuma. 2005. Distribution of Burkholderia cepacia complex species among isolates recovered from persons with or without cystic fibrosis. J. Clin. Microbiol. 43:2926-2928.
46. Riedel, K., C. Arevalo-Ferro, G. Reil, A. Gorg, F. Lottspeich, and L. Eberl. 2003. Analysis of the quorum-sensing regulon of the opportunistic pathogen Burkholderia cepacia H111 by proteomics. Electrophoresis 24:740-750.
47. Rinderknecht, H., M. C. Geokas, P. Silverman, and B. J. Haverback. 1968. A new ultrasensitive method for the determination of proteolytic activity. Clin. Chim. Acta 21:197-203.
48. Rutherford, K., J. Parkhill, J. Crook, T. Horsnell, P. Rice, M. A. Rajandream, and B. Barrell. 2000. Artemis: sequence visualization and annotation. Bioinformatics 16:944-945.
49. Sajjan, U., C. Ackerley, and J. Forstner. 2002. Interaction of cblA/adhesin-positive Burkholderia cepacia with squamous epithelium. Cell. Microbiol. 4:73-86.
50. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
51. Schagger, H., and G. von Jagow. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379.
52. Sokol, P. A., C. Kooi, R. S. Hodges, P. Cachia, and D. E. Woods. 2000. Immunization with a Pseudomonas aeruginosa elastase peptide reduces severity of experimental lung infections due to P. aeruginosa or Burkholderia cepacia. J. Infect. Dis. 181:1682-1692.
53. Sokol, P. A., D. E. Ohman, and B. H. Iglewski. 1979. A more sensitive plate assay for detection of protease production by Pseudomanas aeruginosa. J. Clin. Microbiol. 9:538-540.
54. Sokol, P. A., U. Sajjan, M. B. Visser, S. Gingues, J. Forstner, and C. Kooi. 2003. The CepIR quorum-sensing system contributes to the virulence of Burkholderia cenocepacia respiratory infections. Microbiology 149:3649-3658.
55. Speert, D. P., D. Henry, P. Vandamme, M. Corey, and E. Mahenthiralingam. 2002. Epidemiology of Burkholderia cepacia complex in patients with cystic fibrosis, Canada. Emerg. Infect. Dis. 8:181-187.
56. Sun, L., R. Z. Jiang, S. Steinbach, A. Holmes, C. Campanelli, J. Forstner, U. Sajjan, Y. Tan, M. Riley, and R. Goldstein. 1995. The emergence of a highly transmissible lineage of cbl+ Pseudomonas (Burkholderia) cepacia causing CF centre epidemics in North America and Britain. Nat. Med. 1:661-666.
57. Tomich, M., C. A. Herfst, J. W. Golden, and C. D. Mohr. 2002. Role of flagella in host cell invasion by Burkholderia cepacia. Infect. Immun. 70:1799-1806.
58. Venturi, V., A. Friscina, I. Bertani, G. Devescovi, and C. Aguilar. 2004. Quorum sensing in the Burkholderia cepacia complex. Res. Microbiol. 155:238-244.
59. Visser, M. B., S. Majumdar, E. Hani, and P. A. Sokol. 2004. Importance of the ornibactin and pyochelin siderophore transport systems in Burkholderia cenocepacia lung infections. Infect. Immun. 72:2850-2857.
60. West, S. E., H. P. Schweizer, C. Dall, A. K. Sample, and L. J. Runyen-Janecky. 1994. Construction of improved Escherichia-Pseudomonas shuttle vectors derived from pUC18/19 and sequence of the region required for their replication in Pseudomonas aeruginosa. Gene 148:81-86.(C. Kooi, B. Subsin, R. Ch)