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Journal of Bacteriology, May 2003, p. 3167-3178, Vol. 185, No. 10
0021-9193/03/$08.00+0     DOI: 10.1128/JB.185.10.3167-3178.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Energy-Generating Enzymes of Burkholderia cepacia and Their Interactions with Macrophages

Vasu Punj, Rachna Sharma, Olga Zaborina, and A. M. Chakrabarty^*

Department of Microbiology and Immunology, University of Illinois College of Medicine, Chicago, Illinois 60612

Received 10 January 2003/ Accepted 25 February 2003

Top Abstract Introduction Materials and Methods Results Discussion References

 
We previously demonstrated that several clinical and environmental isolates of Burkholderia cepacia secreted ATP-utilizing enzymes to the medium; the secretion of these enzymes by cystic fibrosis lung isolate strain 38 was shown to be greatly enhanced in the presence of ₂-macroglobulin. Fractionation of the growth medium of cystic fibrosis isolate strain 71 belonging to genomovar I demonstrated the presence of two additional proteins, homologues of Pseudomonas aeruginosa azurin and cytochrome c₅₅₁, which are normally involved in electron transfer during denitrification. A Q-Sepharose column flowthrough fraction of the growth medium of B. cepacia strain 71 enriched with the azurin and cytochrome c₅₅₁ homologues triggered apoptosis in macrophages and mast cells, leading to their death. Incubation of the Q-Sepharose column flowthrough fraction with antiazurin and anti-cytochrome c₅₅₁ antibodies greatly reduced cell death. We cloned and hyperexpressed a gene from B. cepacia strain 71 that encodes the homologue of P. aeruginosa azurin. Such azurin homologues were detected in the growth medium of several strains belonging to genomovars I, III, and VI but not in the growth medium of strains belonging to other genomovars. The growth medium of the strains that elaborated the azurin homologue had high cytotoxicity towards macrophages. Purified azurin homologue was shown to induce apoptosis in macrophages in a caspase-dependent manner and was localized in both the cytosol and nucleus when incubated with or microinjected into macrophages. This is an interesting example of the interaction of a bacterial protein normally involved in cellular energetics with macrophages to effect their cell death.

Top Abstract Introduction Materials and Methods Results Discussion References

 
In the last decade, Burkholderia cepacia has emerged as a major pathogen in the lungs of patients suffering from cystic fibrosis and in patients with chronic granulomatous disease (11, 21, 26, 45). The increasing incidence of infection with B. cepacia in cystic fibrosis patients is reminiscent of similar infections caused by mucoid strains of Pseudomonas aeruginosa (12, 32, 37). However, unlike P. aeruginosa, the population structures of B. cepacia as a group are complex and were divided initially into five genomovars (52), which were further differentiated into new species (27, 53). More recently, polyphasic studies and improved 16S ribosomal DNA-based PCR assays have allowed sensitive and specific identification of various bacterial species, such as Achromobacter xylosoxidans and Ralstonia pickettii in addition to B. cepacia and other previously recognized members of the B. cepacia complex present in the sputum and lungs of cystic fibrosis patients (3, 24). The presence of such hitherto unrecognized bacterial species as well as other Burkholderia species such as B. anthina and B. pyrrocinia (54), has added a degree of complexity to an understanding of the taxonomy, epidemiology, and pathogenesis of the multitude of strains isolated from the lungs of cystic fibrosis patients that are now collectively called B. cepacia complex (28).

Many environmental B. cepacia and Burkholderia sp. strains are efficient biodegraders of toxic chemicals (5, 20, 25, 43, 47) as well as very effective in controlling plant pathogenic soil fungi and nematodes (11, 13). Thus, they have been proposed for environmental release for purposes of toxic chemical bioremediation and enhanced agricultural productivity. Very little, however, is known about the potential pathogenicity of such strains recommended for application in an open environment, leading to the expression of profound concerns about the wisdom of such releases (15, 22). Of particular concern are recent reports that the virulent genomovar III and other strains can be isolated from soil and cannot be easily differentiated from environmental isolates (2, 23).

The virulence factors elaborated by B. cepacia are largely unknown (11), although hemolysins and exopolysaccharides (14, 19, 40) have been implicated in its pathogenicity. We recently demonstrated that, similar to clinical isolates of P. aeruginosa, clinical isolates of B. cepacia secrete a number of ATP-utilizing enzymes that modulate the level of external ATP effluxed from phagocytic cells, leading to their death. Several environmental strains were found to be deficient in the release of these enzymes that acted as potent cytotoxic factors in the presence of millimolar concentrations of ATP. We further demonstrated that secretion of the ATP-utilizing enzymes by a cystic fibrosis isolate, strain 38, of B. cepacia was greatly enhanced in the presence of a mammalian protein such as ₂-macroglobulin (33). Nothing is known about whether the ₂-macroglobulin-mediated enhancement of secretion is unique to this strain or occurs in other clinical or environmental strains or whether secretion of other potential virulence factors may also be modulated by ₂-macroglobulin. It is also not known if B. cepacia secretes ATP-utilizing enzymes that act in an ATP-inducible manner or secretes other cytotoxic factors that may operate by an ATP-independent pathway, as reported for P. aeruginosa (58, 59).

In this paper, we report the secretion of two redox proteins, homologues of P. aeruginosa azurin and cytochrome c₅₅₁, and demonstrate that secretion of the azurin homologue correlates with the cytotoxicity demonstrated by the growth medium of B. cepacia genomovars I, III, and VI. Additionally, we demonstrate the induction of apoptosis in macrophages by purified preparations of the azurin homologue, establishing it as a potential virulence factor.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Bacterial strains and media. B. cepacia strains were grown in several media, including protease peptone yeast extract (PPY; 10 g of peptone, 1.5 g of yeast extract, 10 g of sucrose, 5 g of sodium chloride in 1 liter of water), tryptone beef extract medium (TB; 10 g of Bacto tryptone, 3 g of Bacto beef extract in 1 liter of water), and Luria broth (LB) medium. A list of the strains used in various experiments is shown in Table 1. All cultures were seeded directly from glycerol stocks to avoid variation in phenotypic characteristics as a result of genomic plasticity. All cultures were grown on rotary shakers at 32°C.

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TABLE 1. Bacterial strains used in this study

Column chromatographic fractionation of culture supernatant. B. cepacia strain 71 was grown in TB medium to an A₆₀₀ of 1.1 at 32°C. The cells were centrifuged; the supernatant was filtered through 0.22-µm filters and concentrated with an Amicon YM-10 membrane. This supernatant was fractionated on hydroxyapatite, ATP-agarose, and Q-Sepharose columns as described previously (59). The Q-Sepharose column flowthrough fraction and a portion of each column flowthrough fraction were concentrated, and 2 µg of protein from each sample was used for the cytotoxicity assay. Part of the Q-Sepharose fraction was run on sodium dodecyl sulfate-4 to 20% polyacrylamide gel electrophoresis (SDS-4 to 20% PAGE) gel, followed by transfer to a polyvinylidene difluoride membrane for N-terminal amino acid sequence analysis with an automated Edman AB1477A protein sequencer.

Cloning of azurin homologue of B. cepacia. The azurin-encoding gene (azu) was amplified by PCR with B. cepacia genomic DNA as a template with the primers 5'-GCCAAGCTTATGCTACGTAAACTC-3' (forward) and 5'-GCCCTGCAGCGCGCCCATGAAAAAGCC-3' (reverse). The HindIII and PstI restriction enzyme sites were included within the primer to facilitate cloning. The PCR-amplified product was cloned in vector pUC19. The recombinant plasmid was transformed into Escherichia coli JM109 host cells, which were used for hyperexpression of the B. cepacia azu gene induced in the presence of 1 mM isopropyl-ß-D-thiogalactopyranoside (IPTG). The hyperexpressed azurin was purified from the periplasmic fraction of E. coli according to the method described previously (18).

Macrophage cytotoxicity assay. Macrophages derived from J774 cell lines were grown in RPMI 1640 medium containing L-glutamine, buffered with 10 mM HEPES, and supplemented with 10% fetal bovine serum. Cytotoxicity was determined by measuring the release of lactate dehydrogenase from macrophages as described earlier (39). Cytotoxicity of the azurin homologue in J774 cells was determined with the Cell Titer 96 Aqueous One solution cell proliferation assay (Promega Corporation, Madison, Wis.) as recommended by the manufacturer. Briefly, J774 macrophage cells were plated on 96-well culture plates (10⁴ cells per well) and incubated overnight at 37°C. Subsequently, the cells were treated with the azurin homologue in various doses for various times. Untreated cells and cells treated with Triton X-100 were used as negative and positive controls, respectively. At the end of the incubation, 20 µl of the assay solution containing a tetrazolium compound (MTS; [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)]-2H-tetrazolium) and an electron-coupling reagent (phenazine methosulfate) was added to each well, and the cells were incubated for another 3 h at 37°C in a humidified 5% CO₂ incubator. The amount of soluble formazan produced by cellular reduction of MTS was measured by reading absorbance at 490 nm, as described by Vairano et al. (51).

Antibody production and immunoblot analysis. The antigenic profile of the azurin sequence was analyzed with ABI antigen prediction software (56). Antibodies were raised corresponding to the predicted highly antigenic amino acid sequence CKQFTVNLSHPGNLPKN (amino acids 46 to 52). Immunization of rabbits was carried out according to standard protocols, and the titer of antibodies in the serum was determined by indirect enzyme-linked immunosorbent assay.

Immunoblot analysis was performed by electrotransfer of proteins after SDS-PAGE to a Sequi Blot 0.2-µm polyvinylidene difluoride membrane (Bio-Rad) followed by incubation with primary antibodies. Western blotting was done with anti-rabbit immunoglobulin G labeled with horseradish peroxidase and detected with the ECL system (Amersham Biosciences).

Detection of DNA fragmentation by TUNEL assay. For the terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick-end labeling (TUNEL) assay, the ApoAlert DNA fragmentation assay kit (Clontech, Palo Alto, Calif.) was used, and the assay was performed as recommended by the manufacturer. Briefly, macrophages were seeded at a density of 10⁵ cells per ml on LabTek chamber slides for 2 h. The macrophages were then treated with 200 µg of azurin homologue per ml for different time intervals. Simultaneously, cells were treated with phosphate-buffered saline (PBS) (untreated) or 50 µM benzyloxycarboxyl Val-Ala-Asp fluoromethyl ketone (ZVAD-FMK; Clontech), a cell-permeating general caspase inhibitor. The cells were washed and fixed in 4% paraformaldehyde-PBS and permeabilized with chilled 0.2% Triton X-100-PBS for 5 min on ice. The slides were washed with PBS and equilibrated with equilibration buffer.

The tailing reaction was performed with the ApoAlert DNA fragmentation assay kit. A total of 50 µl of terminal deoxynucleotidyltransferase (TdT) mixture, consisting of 44.5 µl of equilibration buffer, 5 µl of nucleotide mixture, and 0.5 µl of TdT enzyme, was evenly spread on the treated area and incubated in a humid chamber at 37°C for 1 h. The reaction was terminated by incubating the slides with 2x SSC (1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate) for 15 min at room temperature. The cells were then stained with 1 µg of propidium iodide per ml and washed. After the assay, a drop of antifade solution was added, and the treated portion of the slide was covered with a coverslip and the edges were sealed with clear nail polish. Slides were viewed within 2 h under an LSM 510 confocal laser microscope equipped with a 40x objective and a dual filter set for green fluorescence (488 nm) and red fluorescence (568 nm).

Preparation of cytosolic macrophage extracts for caspase assays. The cytosolic extracts were prepared, and caspase-3 and caspase-9 activities were determined in these extracts as described by Zhou et al. (60). Briefly, cells were washed with ice-cold phosphate-buffered saline and lysed in cell lysis buffer (50 mM HEPES [pH 7.4], 0.1% 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate [CHAPS], 5 mM dithiothreitol, 0.1 mM EDTA). After centrifugation at 12,000 x g at 4°C for 10 min, the supernatant (cytosol) was used to determine caspase activities. Caspase-3 activity was determined with the ApoAlert caspase-3 assay kit (Clontech) in accordance with the manufacturer's recommendations, based on spectrophotometric detection of the chromophore p-nitroaniline (pNA) released from the substrate Ac-DEVD-pNA (N-acetyl-Asp-Glu-Val-Asp-p-NO₂-aniline) at 405 nm. For caspase-9, the caspase-9 colorimetric assay kit (Chemicon International, Temecula, Calif.) was used, which involves release and quantitation of pNA from the substrate Ac-LEHD-pNA (N-acetyl-Leu-Glu-His-Asp-p-NO₂-aniline). Specific inhibitors of caspase-3 (DEVD-FMK) and caspase-9 (Ac-LEHD-CHO; Calbiochem-Novabiochem Corp., La Jolla, Calif.) were used to determine the specific activation of the two caspases in cells treated with the azurin homologue.

Subcellular fractionation of macrophages. Macrophages were treated with 200 µg of the azurin homologue per ml for 0, 3, 6, and 12 h. Cytosolic extracts and nuclear extracts were prepared essentially by the procedure of Deveraux et al. (7). Briefly, cells were pelleted by centrifugation after washing with cold buffer A (20 mM HEPES [pH 7.5], 10 mM KCl, 1.5 mM MgCl₂, 1 mM EDTA, 1 mM dithiothreitol, and protease cocktail inhibitor). Subsequently, cells were resuspended in the same buffer, incubated for 20 min on ice, and disrupted by 25 passages through a 26-gauge needle. Cell extract was clarified first by low-speed centrifugation and then by centrifugation at 15,000 x g at 4°C for 30 min.

Intracellular localization of azurin homologue. Macrophages (10⁷ cells) were used for subcellular fractionation after azurin treatment (200 µg/ml) for various time intervals. A total of 20 µg of protein from each time point was loaded on SDS-PAGE gels and blotted on a 0.2-µm polyvinylidene difluoride membrane (Bio-Rad, Hercules, Calif.). The azurin was detected with antiazurin antibody.

Microinjection of azurin homologue in J774 macrophages. Macrophages were cultured overnight on 22-mm glass coverslips coated with collagen adhered to a dish. The azurin homologue labeled red with Alexa fluor 568 (Molecular Probes, Eugene, Oreg.) was microinjected into the cytoplasm of single cells with a computer-controlled microinjector (AIS 2) system. All microinjection experiments were performed with a 0.5-s injection time and 100 hPa of pressure with a Zeiss 200 M microscope. Approximately 25 to 50 cells were injected in each dish. After microinjection, cells were further incubated at 37°C for various times. Cells were then fixed, nuclear DNA was stained blue with 4',6'-diamidino-2-phenylindole (DAPI), and fluorescent images were taken with a Zeiss LSM 510 confocal laser microscope.

Detection of cytochrome c release. The cytosolic extracts were prepared from macrophages either untreated or treated with the azurin homologue for various time periods. These extracts were subjected to SDS-PAGE electrophoresis and transferred to polyvinylidene difluoride membranes, and the Western blot was developed with anti-cytochrome c monoclonal antibody. To determine the release of mitochondrial cytochrome c by confocal microscopy, macrophages were grown on coverslips overnight, treated with 200 µg of azurin homologue per ml, and incubated for different time intervals. The release of cytochrome c was detected by immunofluorescence following the procedure of Pervaiz et al. (36) with minor modifications. Briefly, cells were fixed with methanol-acetone (1:1, vol/vol) and incubated with blocking solution (3% bovine serum albumin) overnight at 4°C. The cells were then incubated for 2 h with mouse monoclonal anti-cytochrome c antibody (clone 6H2.B4; BD Biosciences, San Diego, Calif.). After three washes with PBS-1% fetal bovine serum, cells were exposed to fluorescein isothiocyanate-conjugated anti-mouse immunoglobulin G (Sigma Chemical Co., St. Louis, Mo.). Cells were washed extensively, mounted with antifade Vectashield solution with DAPI, and analyzed by confocal microscopy. Cytosolic cytochrome c (fluorescing green) showed a diffuse staining pattern compared to punctate mitochondrial cytochrome c staining in untreated cells.

Nucleotide sequence accession number. The DNA sequence encoding the azurin homologue has been given GenBank accession number AY238602.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Fractionation of growth medium of cystic fibrosis isolate B. cepacia strain 71 and assay for cytotoxicity of different fractions. We previously reported that several clinical isolates of B. cepacia secreted ATP-utilizing enzymes to the medium that caused macrophage and mast cell death through activation of purinergic receptors (33). To examine if B. cepacia produces cytotoxic agents that operate through an ATP-independent (purinergic receptor-independent) pathway, we fractionated the growth medium (supernatant) of the cystic fibrosis isolate B. cepacia strain 71 (Table 1) by column chromatography. The supernatant growth medium remaining after centrifugation of cells was passed through hydroxyapatite, ATP-agarose, and Q-Sepharose columns, and the flowthrough fractions from each column were assayed for ATP-utilizing enzyme activity and cytotoxicity, as measured by release of lactate dehydrogenase from macrophages and mast cells.

Most of the ATP-utilizing enzymes were removed during column chromatography, leaving the Q-Sepharose flowthrough (QSFT) fraction virtually free of ATP-utilizing enzymes. The cytotoxicity of the supernatant, hydroxyapatite column flowthrough, ATP-agarose column flowthrough, and QSFT fractions to J774 cell line-derived macrophages in the absence and in the presence of 1.0 mM ATP is shown in Table 2. At 1.0 mM, ATP itself had substantial cytotoxicity (22%), as reported previously (39). The supernatant, hydroxyapatite flowthrough, and ATP-agarose column flowthrough fractions had low cytotoxicity by themselves (in the absence of ATP), but the cytotoxicities were significantly higher in the presence of ATP. The QSFT fraction, on the other hand, had high cytotoxicity, which could not be further enhanced in the presence of ATP (Table 2). Thus, the growth medium of B. cepacia strain 71 had ATP-independent cytotoxic activity as well.

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TABLE 2. Cytotoxicity of supernatant and various flowthrough chromatographic fractions from hydroxyapatite, ATP-agarose, and Q-Sepharose columnsa

To see if the ATP-independent cytotoxicity was limited to macrophages or might also affect other phagocytic cells, we examined the cytotoxicity of the strain 71 QSFT fraction towards mast cells. The supernatant, hydroxyapatite flowthrough, and ATP-agarose flowthrough fractions demonstrated low cytotoxicity, while the QSFT fraction showed high cytotoxicity in the absence of ATP (data not shown), confirming that the cytotoxic agent(s) present in the QSFT fraction is active against mast cells as well.

Presence of homologues of azurin and cytochrome c₅₅₁ in QSFT fraction. In order to determine the nature of the proteins present in the QSFT fraction that might be responsible for the death of macrophages and mast cells, we ran the supernatant, ATP-agarose flowthrough, hydroxyapatite flowthrough, and QSFT fractions on SDS-PAGE. While the supernatant, ATP-agarose flowthrough, and hydroxyapatite flowthrough fractions had multiple protein bands, the QSFT fraction had three major protein bands of 8 kDa, 21 kDa, and 75 kDa (data not shown). N-terminal amino acid sequencing of the 8-kDa (EDPEVLFKNK) and the 21-kDa (AXXSVDIQGN, where X is a residue of uncertain identity) bands showed 100% and 80% sequence identity with that of P. aeruginosa cytochrome c₅₅₁ and the copper-containing redox protein azurin, respectively (55). We have yet to characterize the 75-kDa protein.

To analyze whether the azurin and cytochrome c₅₅₁ homologues of B. cepacia were indeed responsible for the death caused by the QSFT fraction, we determined if a mixture of antiazurin and anti-cytochrome c₅₅₁ (2 µg of each) antibodies would have any effect on the cytotoxicity exhibited by the B. cepacia QSFT fraction. Polyclonal antibodies against purified azurin and cytochrome c₅₅₁ were prepared for this purpose. When a mixture of azurin and cytochrome c₅₅₁ was tested for cytotoxicity against J774 cell line-derived macrophages, high cytotoxicity was observed (Fig. 1, column A+C). When this mixture was incubated with a mixture of antiazurin and anti-cytochrome c₅₅₁ antibodies (4 µg of protein) and then tested for macrophage cytotoxicity, the cytotoxicity was greatly diminished (Fig. 1, column A+C+Ab4), suggesting that antibody treatment neutralizes the cytotoxicity mediated by azurin plus cytochrome c₅₅₁.

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FIG. 1. Neutralization of macrophage cytotoxicity by a mixture of antiazurin and anti-cytochrome c551 antibodies. Macrophage cytotoxicity was determined by the lactate dehydrogenase (LDH) release assay. A mixture of azurin and cytochrome c551 (A+C) was used as a positive control. The cytotoxicity associated with this combination was neutralized by treatment with antibodies against azurin and cytochrome c551 (A+C+Ab 4). When macrophages treated with various dilutions of antibody mixtures (Ab 0.5 to Ab 4) were treated with 2 µg of the QSFT fraction, a gradual decrease in cytotoxicity was observed with increasing concentrations of antibody mixture. The numbers after Ab represent micrograms of protein. Incubation with preimmune serum, even at high concentrations, had no effect on cytotoxicity (data not shown).

When the B. cepacia QSFT fraction was tested for cytotoxicity, high cytotoxicity was also observed (Fig. 1, column QSFT). Pretreatment of the QSFT fraction with increasing amounts (0.5 to 4 µg of protein) of antiazurin and anti-cytochrome c₅₅₁ antibodies greatly reduced this cytotoxicity (Fig. 1, columns QSFT+Ab 0.5 to Ab 4), demonstrating a major role of azurin and cytochrome c₅₅₁ in the QSFT fraction for imparting cytotoxicity. Treatment with increasing amounts (0.5 to 4 µg of protein) of preimmune serum, however, did not lead to appreciable loss of cytotoxicity (data not shown), confirming that the antiazurin and anti-cytochrome c₅₅₁ antibodies led to neutralization of homologues of such proteins in the QSFT fraction, thereby contributing to the loss of cytotoxicity. We have not yet determined the role played by cytochrome c in B. cepacia virulence. The cloning of its gene and its hyperexpression are currently under investigation.

Cloning and hyperexpression of gene encoding B. cepacia azurin homologue. While the loss of cytotoxicity of the QSFT fraction on treatment with antiazurin plus anti-cytochrome c₅₅₁ antibodies provided insights into the potential role of these redox proteins in cytotoxicity, it was necessary to test the involvement of the azurin and cytochrome c₅₅₁ homologues in B. cepacia virulence directly. As a first step towards this goal, we cloned the azu gene encoding the azurin homologue from B. cepacia as described under Materials and Methods. The amino acid sequence of the B. cepacia strain 71 (genomovar I) azu gene product showed significant sequence identity with the azurin sequences of a number of other bacteria (Fig. 2), including some, such as Achromobacter xylosoxidans, which can also be recovered from the sputum of cystic fibrosis patients (24).

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FIG. 2. Multiple alignment of B. cepacia azurin protein homologue sequence with sequences of azurin from various organisms. Clustal W software was used to generate the alignment. *, amino acids conserved among all the proteins; , amino acids conserved in at least three of six organisms; #, nonconserved amino acids (50). The signal sequence (amino acids 1 to 18) is highly variable among all the proteins.

We explored if the azurin homologue could be detected in the growth medium of B. cepacia strain 71 during growth in L broth. Western blotting data demonstrated the presence of the azurin homologue in the growth medium filtered through a 0.22-µm filter during early to mid-log phase (data not shown). Then we explored if the azurin homologue could be detected in the filtered growth medium of various B. cepacia genomovars, as depicted in Table 1. Of the nine genomovars tested, only strain 71 (genomovar I), strain PC783 (genomovar I), strain HI2147 (genomovar III), and strain AU0649 (genomovar VI) demonstrated the presence of an azurin homologue in the growth medium (Fig. 3A). All the other strains were negative in this respect.

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FIG. 3. Release of azurin homologue during growth of various strains of B. cepacia complex in L broth as detected by Western blotting. Even though all the strains (Table 1) were tested, only those that released the azurin homologue are shown (A). These strains also showed an enhanced level of azurin in the growth medium when they were grown in the presence of 2-macroglobulin (1 mg/ml) (B). All the strains were grown in the presence and absence of 2-macroglobulin to an optical density at 600 nm of 1.1. The cells were then centrifuged, supernatants were filtered through a 0.22-µm filter, and high-molecular-weight proteins were separated by passage through Centricon YM100 (Amicon) and precipitated with ammonium sulfate. Excess salt was removed by extensive washing in Centricon YM10, and then 30 µg of proteins was separated on SDS-PAGE and blotted with antiazurin antibodies to detect the presence of the azurin homologue as described in Materials and Methods. The amount of azurin was quantified with NucleoTech Gel Expert 97 software.

Similar to the ATP-utilizing enzymes whose secretion was stimulated in the presence of 1 mg of ₂-macroglobulin per ml in the growth medium (33), the secretion of the azurin homologue was stimulated to various degrees during growth of the four strains in the presence of 1 mg of ₂-macroglobulin per ml (Fig. 3B). When the growth medium (5 µg of protein each) from the above strains was examined for cytotoxicity towards macrophages, significant cytotoxicity was observed only with the growth medium of the strains that secreted azurin homologues in significant amounts (Table 3). The high cytotoxicity thus reflects a high level of secretion of azurin when the cells were grown in the presence of ₂-macroglobulin.

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TABLE 3. Cytotoxicity of supernatants of various strains grown in the presence of 2-macroglobulina

B. cepacia azurin homologue induces apoptosis in macrophages. Since our primary goal was to study the interaction of redox proteins such as azurin with macrophages, it was important for us to evaluate the role of the B. cepacia azurin homologue in macrophage cytotoxicity. The cytotoxicity was determined by the MTS assay as described under Materials and Methods. Recombinant B. cepacia azurin homologue demonstrated high cytotoxicity towards J774 cell line-derived macrophages, and this cytotoxicity was concentration dependent (Fig. 4A). When the azurin homologue was used at 200 µg/ml for various time periods, the maximal cytotoxicity was reached between 24 and 32 h (Fig. 4B).

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FIG. 4. Cytotoxicity of the azurin homologue of the B. cepacia towards J774 cells as determined by MTS assay as described in Materials and Methods. (A) Cytotoxicity of macrophages treated for 24 h with various concentrations of the azurin homologue. (B) Cytotoxicity of macrophages treated with 200 µg of the azurin homologue per ml for various times.

To define the nature of the macrophage cell cytotoxicity, we tested to see if the purified azurin homologue could cause cell death through the induction of apoptosis. One of the hallmarks of apoptosis is DNA fragmentation; we used the TUNEL assay (8) to determine the extent of incorporation of fluorescently tagged fluorescein-dUTP (Clontech ApoAlert DNA fragmentation assay kit) into the nuclear DNA of untreated and azurin-treated macrophages. Extensive dUTP incorporation occurs only in apoptotic cells with considerable nuclear DNA fragmentation, giving rise to cells in which the nuclear DNA fluoresces green. Untreated macrophages did not show any significant nuclear green fluorescence when incubated with fluorescein-dUTP in the TUNEL assay (Fig. 5A, untreated), showing little dUTP incorporation in nuclear DNA. When macrophages were treated with the B. cepacia azurin homologue for 6 h, 12 h, and 16 h, increasing green fluorescence-tagged macrophages were seen (Fig. 5A, as indicated) under the green channel (bottom, left); the red channel (top, left) showed all macrophages, both healthy and apoptotic. The superimpositions of the red and green channels showed macrophages with those incorporating dUTP in their DNA as yellow (lower right-hand channels in Fig. 5A). When macrophages were treated with the azurin homologue in the presence of the general caspase inhibitor ZVAD-FMK, very few cells were seen to undergo apoptotic changes (Fig. 5B).

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FIG. 5. TUNEL assay for detection of apoptosis-induced nuclear DNA fragmentation in azurin homologue-treated macrophages. The assay is based on terminal deoxynucleotidyltransferase-mediated dUTP nick-end labeling, where Tdt catalyzes the incorporation of fluorescein-dUTP at the free 3'-hydroxyl ends of fragmented DNA in cells undergoing apoptosis. The incorporation of fluorescein-dUTP into the fragmented nuclear DNA generates the green fluorescence detected by confocal microscopy. (A) J774 cell-line derived macrophages were grown on LabTek chamber slides and incubated for 6, 12, and 16 h. A negative control (untreated) without azurin treatment (treated with TM buffer for 16 h) was also maintained. Macrophages viewed under both red and green channels are shown. (B) Macrophages treated with azurin homologue (16 h) and 50 µM general caspase inhibitor ZVAD-FMK, showing inhibition of DNA fragmentation induced by the azurin homologue.

One of the mitochondrial events associated with apoptosis is an alteration in the mitochondrial membrane potential, leading to cytosolic translocation of cytochrome c from the intermembrane space of mitochondria (1). We therefore measured the release of macrophage mitochondrial cytochrome c to the cytosol by confocal microscopy. In macrophages treated with buffer (untreated) or after 6 h of treatment with the azurin homologue (200 µg/ml), most of the cytochrome c was found as granules in the mitochondria showing a punctate staining pattern (Fig. 6A). After 12 h of treatment, however, the cytochrome c was found throughout the cytosol, showing diffuse staining in a few macrophages undergoing apoptosis (Fig. 6A). When the cytosolic fractions (30 µg of protein each) free of mitochondria were examined with anti-cytochrome c antibodies by Western blotting, very little cytochrome c was detected in the cytosol after 3 or 6 h of treatment with the azurin homologue, but some accumulation was observed after 12 h of treatment (Fig. 6B). When the level of ß-actin, a cytosolic protein, was examined as an internal control by Western blotting, its level remained fairly constant throughout the treatment process, suggesting that 12 h of treatment of macrophages with the azurin homologue elicited the release of mitochondrial cytochrome c to the cytosol before the onset of apoptosis mediated by the azurin homologue.

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FIG. 6. Time course of mitochondrial cytochrome c release into the cytosol of macrophages during treatment with the azurin homologue. (A) Macrophages were grown on coverslips and treated with the azurin homologue (200 µg/ml) for the indicated times; mitochondrial cytochrome c localization was determined by confocal microscopy with anti-cytochrome c antibody as described in Materials and Methods. The nucleus was stained blue with DAPI. (B) Cytosolic extracts of azurin homologue-treated and untreated (0 h) macrophages taken at the times indicated were separated by SDS-PAGE, transferred to a polyvinylidene difluoride membrane, and subjected to Western blot analysis with mitochondrial anti-cytochrome c (Cyt C) antibody.

Since release of mitochondrial cytochrome c to the cytosol triggers caspase cascade activation (1), we assayed the level of two members of the caspase cascade, caspase-3 and caspase-9, that are known to be activated during apoptosis due to release of mitochondrial cytochrome c (42, 46). Macrophages were incubated for various time periods in the absence or in the presence of the azurin homologue (200 µg/ml) and washed thoroughly, and cytosolic extracts were prepared as described in Materials and Methods. Caspase-3 activity was assayed with 60 µg of cytosolic protein extract with the colorimetric substrate Ac-DEVD-pNA at 37°C for 1 h. Caspase-9 was assayed similarly with Ac-LEHD-pNA as a substrate. Specific inhibitors of both caspase-3 (Ac-DEVD-FMK) and caspase-9 (Ac-LEHD-CHO) were also included to determine the specificity of the caspase reactions. The results clearly demonstrated that both caspase-3 and caspase-9 (Table 4) activities increased with time in macrophage cytosolic extract when the macrophages were treated with the azurin homologue. The caspase activities remained low when the macrophages were treated with buffer (untreated) and when the extracts from treated macrophages were assayed in the presence of caspase inhibitors.

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TABLE 4. Measurement of caspase activities in cytosolic extracts of macrophages treated or not with the azurin homologuea

Trafficking of azurin homologue to macrophage nucleus. Since incubation of macrophages with the purified azurin homologue protein triggered apoptosis, it is clear that the azurin homologue either exerts its effect as a surface-bound protein or enters the cell cytosol. The role of any macrophage surface receptor(s) in the internalization of the azurin homologue is unknown at present. To evaluate the localization of the azurin homologue in J774 cell line-derived macrophages, we incubated the macrophages for 0, 3, 6, and 12 h, washed them, made extracts, and isolated various subcellular fractions. Western blotting was then performed with antiazurin antibody with 20 µg of protein from each fraction. We also microinjected the azurin homologue into the macrophage cells and used confocal microscopy to detect the presence of azurin in the cytosolic and nuclear fractions.

When the purified azurin homologue was incubated with the macrophages and subcellular fractions (cytosol and nuclear) were examined, azurin was found (Fig. 7A) in the cytosolic fraction both during early (3 h) and later (6 h and 12 h) periods of incubation. In contrast, very little azurin homologue was found in the nuclear fraction at 3 h but was detected in large amounts at 6 h and 12 h of incubation (Fig. 7A). We also microinjected the azurin homologue directly into the macrophage cytosol to examine its nuclear trafficking. Confocal microscopy of the microinjected macrophages confirmed the presence of the azurin homologue in the cytosol at 15 min; by 3 h, azurin was also found in the nucleus (Fig. 7B), suggesting that the cytosolic azurin homologue can traffic to the nucleus during incubation of the macrophages with purified protein or after microinjection of the azurin homologue into the macrophage cytosol.

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FIG. 7. Subcellular localization and trafficking of the azurin homologue. (A) Localization of the azurin homologue in the cytosol and nuclear fractions of macrophages treated with 200 µg of the protein per ml for the indicated times was determined by Western blotting with antiazurin antibodies. (B) Representative confocal microscopy images for trafficking of the azurin homologue from the cytosol to the nucleus of macrophages. Microinjection of Alexa Fluor 568-labeled azurin homologue is described in Materials and Methods. After microinjection, the macrophage cells were incubated for the indicated time periods. The insets show magnified images of single injected cells after 15 min and 3 h. Azurin is labeled red (Alexa Fluor 568), while the nucleus is stained blue with DAPI. The arrows indicate the entry of the azurin homologue in the nucleus.

Top Abstract Introduction Materials and Methods Results Discussion References

 
One of the goals of this study was to understand how bacterial enzymes that are normally involved in cellular energetics interact with mammalian cells such as macrophages, ultimately triggering their death. The role of ATP-utilizing enzymes in the induction of macrophage cell death has been reported previously for both P. aeruginosa (58) and B. cepacia (33). Both the Pseudomonas and Burkholderia genera are well known for their nutritional versatility as well as for their ability to cause fatal infections in the lungs of cystic fibrosis patients, yet the nature and outcome of such infections are often different. P. aeruginosa is an extracellular pathogen that causes chronic infections in cystic fibrosis patients; in contrast, B. cepacia is known to be capable of some intracellular replication (30, 31, 41) and causes rapid morbidity and mortality in such patients. This suggests that the nature of the virulence factors or the way in which they act must be different.

Both P. aeruginosa and B. cepacia elaborate ATP-utilizing enzymes that have been reported to cause macrophage cell death; however, it is not known if they act similarly. For example, ADP-ribosylating enzymes and toxins of P. aeruginosa such as exotoxin A and exoenzyme S, as well as cholera toxin produced by Vibrio cholerae, even though they have the common ADP-ribosylating activity, all have different targets and different modes of action.

We have reported that P. aeruginosa elaborates azurin and cytochrome c into the growth medium that induce cell death through complex formation with and stabilization of tumor suppressor protein p53 (57). The azurin homologue of B. cepacia has now been shown to induce apoptosis in macrophages. Given the differences in the mode of action of these two pathogens, it would be interesting to know if both of them have the same mode of action. An interesting example in this regard is the elaboration of a pore-forming protein, listeriolysin O, secreted by the intracellular pathogen Listeria monocytogenes and a related pore-forming protein, perfringolysin O, secreted by the extracellular pathogen Clostridium perfringens. The presence of listeriolysin O in L. monocytogenes allows the bacterium to escape from the macrophage vacuole to reach the macrophage cytosol and live there but does not allow lysis of the plasma membrane of the macrophage and its killing. Lysteriolysin O and perfringolysin O have 43% sequence identity and 70% sequence similarity. While highly homologous, they have very different modes of action, since extracellular C. perfringens does not need to live in the macrophage and therefore perfringolysin O allows lysis of the macrophage plasma membrane, leading to the death of the cell. A small addition of 27 amino acids with a PEST-like sequence in the L. monocytogenes protein accounts for this difference, since transfer of this sequence to perfringolysin O transformed that toxic cytolysin into a nontoxic derivative that facilitated intracellular growth (6). It would be of great interest to know if the P. aeruginosa azurin and B. cepacia azurin homologues exhibit differences in their mechanism of cytotoxicity towards macrophages.

We have reported that the secretion of ATP-utilizing enzymes by B. cepacia strain 38 and others is greatly enhanced in the presence of ₂-macroglobulin (33). It is interesting that the amount of the azurin homologue present in the growth medium of B. cepacia strains (Fig. 3A) belonging to genomovars I, III, and VI is also greatly increased in the presence of ₂-macroglobulin (Fig. 3B), and there is a correlation between the elaboration of the azurin homologue in the growth medium and the amount of cytotoxicity exhibited by that growth medium (Table 3).

Even though both the ATP-utilizing enzymes and the azurin homologue respond to a common mammalian protein such as ₂-macroglobulin, which may enhance the secretory mechanism of both of these virulence factors, the mechanism of cell killing by these factors is very different. The ATP-utilizing enzymes are active mostly in the presence of ATP, operating through activation of the P2Z purinergic receptors (58). Indeed, similar modulation of purinergic receptor activation by secreted ATP-utilizing enzymes of the parasitic nematode Trichinella spiralis has recently been reported (9, 10). In contrast, the azurin homologue triggers the release of mitochondrial cytochrome c to the cytosol (Fig. 6), resulting in elevated caspase-3 and caspase-9 levels (Table 4) and nuclear DNA fragmentation (Fig. 5). Further details of the mode of action of the B. cepacia homologue are currently under investigation.

One of the most intriguing and difficult aspects of B. cepacia infection in the lungs of cystic fibrosis patients is the diversity of various B. cepacia-like organisms, as described in Table 1. It is by no means clear which microorganisms are truly virulent infective agents and which are simply commensal carryovers or even hardy contaminants growing in a somewhat weakened immune system. We have examined only a single strain from each genomovar (two strains from genomovar I), which is totally inadequate to allow us to draw any general conclusions. Nevertheless, our data imply some degree of correlation between the azurin-mediated cytotoxicity exhibited by members of genomovars I, III, and VI and the frequency of isolation of such strains from the lungs of cystic fibrosis patients, particularly members of genomovar III. Many more members of the B. cepacia complex assigned to different genomovars need to be examined for the ability to secrete the azurin homologue and for severity of infection before any meaningful conclusions can be drawn about azurin's potential as a virulence factor of the B. cepacia complex.

Finally, one may wonder why ATP-utilizing enzymes and redox proteins elaborated by the pathogens found in the cystic fibrosis lung, B. cepacia and P. aeruginosa, that are normally involved in the energetics of the cell are also involved in mammalian cell death. It is interesting that these pathogens preferably release these enzymes in response to a mammalian host protein such as ₂-macroglobulin or -casein (33, 58). An analogy to this is the release from mitochondria of similar enzymes such as cytochrome c and the apoptosis-inducing factor AIF, an oxidoreductase flavoprotein, in the presence of death signals such as withdrawal of growth factors, presence of DNA-damaging agents, chemotherapeutics, etc., leading to cell death (1, 4, 35).

It is also interesting that adenylate kinase is secreted by P. aeruginosa as a virulence factor (29) and is surface exposed in Streptococcus agalactiae, the causative agent of sepsis, pneumonia, and meningitis in neonates (49), but is also released simultaneously with cytochrome c from the intermembrane space of mitochondria during apoptosis (16, 44). Mitochondria are the storehouse of the energetics of eukaryotes, harboring both the electron transport chain and the machinery for ATP synthesis. Mitochondria, of course, are prokaryote-like structures which are believed to have evolved hundreds of millions of years ago when the ancestral eukaryotic cells entered into a mutually beneficial partnership with the ancestors of the present-day bacteria that allowed the eukaryotic cells to utilize the energy-generating machinery of the prokaryotes to take advantage of moving from an anaerobic to an increasingly oxygen-rich environment (17, 34). Unlike some obligate endosymbiotic bacteria of aphids, in which the genome has remained fairly stable for the past 50 to 70 million years (48), the prokaryotic ancestors of the protomitochondria eventually lost many of their essential genes, transferring some to the eukaryotic nucleus and thereby becoming an obligate endosymbiotic organelle.

Since mitochondria are central to mammalian cell apoptosis, in which release of AIF or cytochrome c plays an important role, it appears that present-day prokaryotes such as B. cepacia, P. aeruginosa, and presumably others retain the ability to use their energy-generating machinery in the form of ATP-utilizing enzymes or redox proteins to effect mammalian cell death, much to their advantage in coping with a nonsymbiotic hostile environment (38). The present study thus provides additional evidence of the interesting role of bacterial proteins that are normally involved in cellular energetics in mammalian cell death.

 
This work was supported by PHS grant ES04050-17 from the National Institute of Environmental Health Sciences.

We thank J. LiPuma and E. Mahenthiralingam for providing the strains mentioned in Table 1.

 
* Corresponding author. Mailing address: Dept. of Microbiology and Immunology, University of Illinois College of Medicine, 835 South Wolcott Avenue, Chicago, IL 60612. Phone: (312) 996-4586. Fax: (312) 996-6415. E-mail: pseudomo{at}uic.edu.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Journal of Bacteriology, May 2003, p. 3167-3178, Vol. 185, No. 10
0021-9193/03/$08.00+0     DOI: 10.1128/JB.185.10.3167-3178.2003
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