1 Robert Koch-Institut, Nordufer 20, 13353 Berlin, Germany
2 Lehrstuhl für Mikrobiologie, Friedrich-Alexander-Universität Erlangen-Nürnberg, Staudtstr. 5, 91058 Erlangen, Germany
3 Department of Microbiology, University of Alabama at Birmingham, Birmingham, AL 35294, USA
Correspondence
Astrid Lewin
Lewina{at}rki.de
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ABSTRACT |
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INTRODUCTION |
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M. smegmatis belongs to the fast-growing opportunistic mycobacteria. Although M. smegmatis is generally considered to be an environmental saprophytic bacterium, it can cause skin and soft-tissue lesions (Brown-Elliott & Wallace, 2002). Lung infections caused by M. smegmatis occur rarely (Daley & Griffith, 2002
; Howard & Byrd, 2000
; Kumar et al., 1998
; Schreiber et al., 2001
; Vonmoos et al., 1986
). However, M. smegmatis has been identified as a causative agent of fatal disseminated disease in patients with IFN-
receptor deficiencies (Andrews & Sullivan, 2003
; Howard & Byrd, 2000
; Jouanguy et al., 1999
; Pierre-Audigier et al., 1997
).
An interesting feature of many mycobacterial species is their ability to survive inside amoebae, leading to the classification of mycobacteria as amoeba-resistant micro-organisms' (Greub & Raoult, 2004). The mechanisms used by macrophages and amoebae for phagocytosis, phagolysosome formation and digestion of intracellular bacteria are very similar (Allen & Dawidowicz, 1990a
, b
; Brown & Barker, 1999
; Greub & Raoult, 2004
; Winiecka-Krusnell & Linder, 2001
). Reciprocally, the strategies employed by bacteria to escape destruction by macrophages or amoebae are also similar. This supports the theory that an evolutionary selection for survival in environmental protozoa has enabled intracellular pathogenic bacteria to develop the capacities necessary for survival in macrophages (Brown & Barker, 1999
; Steinert et al., 1998
; Winiecka-Krusnell & Linder, 2001
). In this context, it is interesting that passage through amoebae can enhance the virulence of pathogenic intracellular bacteria and that there exists a correlation between the virulence of mycobacterial species and survival in amoebae (Cirillo et al., 1997
).
All mycobacteria are characterized by a thick hydrophobic cell wall, which is penetrated by porins mediating diffusion of small hydrophilic nutrients into the cell. MspA from M. smegmatis belongs to a novel class of porins present in many fast-growing mycobacteria but apparently absent in slow-growers (Niederweis et al., 1999). MspA is an extremely stable octameric protein composed of 20 kDa monomers (Faller et al., 2004
; Heinz et al., 2003
). In addition to the mspA gene, M. smegmatis possesses three homologous genes named mspB, mspC and mspD. The main diffusion pathway of M. smegmatis is provided by MspA (Engelhardt et al., 2002
; Stahl et al., 2001
).
In a previous study, we showed that the intracellular persistence of a M. bovis BCG derivative expressing the mspA gene from M. smegmatis was enhanced (Sharbati-Tehrani et al., 2004), indicating that the cell wall permeability may directly determine intracellular survival of mycobacteria. To further address this question and analyse the effects of cell wall permeability on intracellular persistence, M. smegmatis was chosen, since porin genes from this species have been intensively characterized and mutagenized (Stahl et al., 2001
; Stephan et al., 2004b
).
To find out if porins from M. smegmatis influence the outcome of the infection process, we used two porin mutants from M. smegmatis and analysed their extracellular and intracellular growth. In the first mutant, mspA was deleted (Stahl et al., 2001), while in the second mutant, mspA and mspC were deleted (Stephan et al., 2004b
). The intracellular persistence of the mutants compared to the parental strain was analysed in different phagocytic cells, including amoebae. We demonstrate that the presence of porins reduces the ability of M. smegmatis to survive in vivo.
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METHODS |
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The plasmid pMN013 was used for complementation of porin mutations in the strains MN01 and ML10 and carries a transcriptional fusion of the promoter pimyc with the mspA gene (Mailaender et al., 2004). Plasmids were introduced into M. smegmatis derivatives via electroporation, as described previously for M. bovis BCG (Sharbati-Tehrani et al., 2004
).
Cell lines and culture conditions.
Axenic Acanthamoeba castellanii (Walochnik et al., 2000) was grown to 90 % confluence at 28 °C in the dark in BD Falcon 75 ml flasks (BD Biosciences) containing PYG broth (Moffat & Tompkins, 1992
). The mouse macrophage cell line J774A.1 (DSMZ no. ACC170) was maintained as previously described (Lewin et al., 2003
). Murine bone marrow macrophages (BMMs), kindly provided by Stefan Kaulfuss, were derived in vitro from bone marrow progenitors of black female C57BL/6 mice, as described previously (Dorner et al., 2002
). Prior to infection, BMMs were maintained in D-MEM (Gibco) supplemented with 10 % fetal calf serum (Biochrom AG), 5 % horse serum (Biochrom AG), 1 % 1 mM sodium pyruvate and 1 % L-glutamine.
Growth experiments in broth.
Determination of growth of M. smegmatis strains SMR5, MN01 and ML10 in vitro was carried out in Middlebrook 7H9 medium at pH 5·0 and pH 6·7. Prior to inoculation, exponential-phase M. smegmatis cells were washed twice with PBS supplemented with 0·05 % Tween 80 (PBS-T) to minimize the formation of aggregates. Afterwards, 120 ml of medium was inoculated with 3x107 c.f.u., evenly distributed into three flasks and incubated at 37 °C without shaking. Growth was determined by measuring the OD600 of cultures in triplicate.
Infection of macrophages and measurement of intracellular growth.
Infection of the macrophage cell line J774A.1 and of the BMMs was performed as previously described (Sharbati-Tehrani et al., 2004), with the following modifications: 5x104 cells per well were seeded in 24-well plates (BD Biosciences); J774A.1 cells were allowed to adhere for 2 h; BMMs were allowed to adhere overnight. Cells were then infected with exponential-phase M. smegmatis strains at an m.o.i. of 1 in triplicate. After 4 h the supernatants were removed and adherent cells were washed twice with medium. The cells were then treated with 200 µg amikacin ml1 for 1 h to kill the non-phagocytosed M. smegmatis. After washing twice with medium, 1 ml medium supplemented with 2 µg amikacin ml1 was added to each well to prevent extracellular growth. Samples for quantification of intracellular bacteria were taken at the end of the infection period after removal and killing of extracellular bacteria and then twice per day until 54 h post-infection. After removal of supernatants, lysis of cells was performed by the addition of 1 ml sterile distilled H2O and incubation at 37 °C until lysis was complete. The intracellular persistence of M. smegmatis was determined by plating and colony counting.
Infection of A. castellanii.
Prior to infection, A. castellanii monolayers were washed with A. castellanii buffer (Moffat & Tompkins, 1992) and then harvested and resuspended in A. castellanii buffer. Then 105 A. castellanii per well were seeded in 24-well plates (BD Biosciences) and allowed to adhere for 1 h. Afterwards, amoebae were infected with exponential-phase M. smegmatis strains at an m.o.i. of 10 (Cirillo et al., 1997
) in triplicate. After an initial infection time of 2 h, further treatment was performed according to the infection procedure for J774A.1 cells, replacing the medium with A. castellanii buffer, except that no amikacin was added to A. castellanii buffer after the washing procedure. Intracellular mycobacteria were recovered by lysing the amoebae with PBS supplemented with 0·5 % SDS (Cirillo et al., 1997
). The intracellular persistence of M. smegmatis was determined by plating and colony counting.
Quantification of porin gene expression.
Expression of porin genes in the different strains was determined by means of RT-real-time PCR using the Mx3000P Real-time PCR System (Stratagene). M. smegmatis derivatives were grown to an OD600 of 0·8 and RNA was extracted according to the method of Bashyam & Tyagi (1994). One microgram of the RNA was treated prior to RT-real-time PCR with RQ1 RNase-Free DNase (Promega GmbH). We quantified mspA expression by amplifying a fragment of 100 bp using the primers mspATaqFW (5'-CGTGCAGCAGTGGGACACCTT-3'), mspATaqBW (5'-CCACGATGTACTTGGCGCGAC-3') and the dual-labelled detector probe MspATaqProbe (5'-FAM-TGGACCGCAACCGTCTTACCCGTGAGTG-TAMRA-3'). The reaction was performed with the Access RT-PCR System (Promega) in a 50 µl reaction mix containing 1 µl (100 ng) Dnase-treated RNA as template, 0·2 mM each dNTP, 1 mM MgSO4, 40 pmol each primer, 50 nM probe, 5 U AMV Reverse Transcriptase, 5 U Tfl DNA Polymerase, AMV/Tfl Reaction Buffer and 30 nM ROX as passive reference dye. Amplification was carried out by running a first reverse transcription step at 48 °C for 45 min, followed by 2 min at 94 °C and 40 cycles with 30 s at 94 °C and 1 min at 58 °C. RNA amounts were determined by three measurements for each sample using a calibration curve established with known amounts of linearized pSSa100, which carries one copy of mspA (Sharbati-Tehrani et al., 2004
). Non-reverse-transcribed PCR controls were performed with the same samples to guarantee the absence of contaminating genomic DNA.
Electron microscopy.
For transmission electron microscopy (TEM) of uninfected and infected A. castellanii, the A. castellanii buffer was replaced by glutaraldehyde (2·5 %, v/v) buffered with HEPES (0·05 M, pH 7·2) and the amoebae were fixed for 1 h at room temperature, then stored at 4 °C in the same solution. The cells were first agarose-block embedded by mixing equal volumes of cells and low-melting-point agarose [3 % in double-distilled H2O (ddH2O)], postfixed with OsO4 for 1 h (1 % in ddH2O; Plano) and block-stained with uranyl acetate for 1 h (2 % in ddH2O; Merck). The samples were then dehydrated stepwise in graded alcohol and embedded in LR-White resin (Science Services), which was polymerized at 60 °C overnight. Ultrathin sections were prepared with an ultramicrotome (Ultracut S, Leica) and placed on naked 400-mesh grids. The sections were stained with lead citrate and stabilized with approximately 1·5 nm carbon (carbon evaporation; BAE 250, Bal Tec). TEM was performed with an EM 902 (Zeiss) using a slow-scan CCD camera (Proscan, Scheuring).
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RESULTS |
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To prove that the enhanced persistence of the mutants was in fact caused by the deletion of the porin genes, we performed complementation experiments by transforming the mutants with a plasmid carrying the mspA gene. The plasmid pMN013 (Mailaender et al., 2004) carries the mspA gene fused to the M. smegmatis promoter pimyc on a shuttle vector. The intracellular persistence of the mutants carrying pMN013 was tested in A. castellanii, because the differences among the strains were most pronounced in this test system. RT-real-time PCR experiments confirmed that complementation of the porin deletion in mutant strains was achieved by introducing pMN013. The complementation of porin deletions by pMN013 resulted in a higher mspA expression in both mutant strains than in the parental strain (Table 2
). Transcription of mspA in the mutants carrying pMN013 was about 710 times stronger than in the wild-type. The shuttle plasmid pMN013 is present in several copies in mycobacteria and the promoter pimyc is a relatively strong promoter (Kaps et al., 2001
). The expression of mspA using pMN013 affected the adaptation of the mutants to intracellular survival, so that they resembled the phenotype of the wild-type SMR5. At 44 h post-infection, 1·1±0·6 % of MN01 (pMN013) and 1·8±0·3 % of ML10 (pMN013) remained intracellular and viable (relative to the 4 h values). The decrease of the intracellular persistence of the mutants harbouring pMN013 confirmed that the deletion of the porin genes was responsible for the enhanced persistence of the mutants in the amoeba.
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DISCUSSION |
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As the enhanced survival of the porin mutants occurred in all three phagocytic systems tested, we wondered which bactericidal mechanism common to macrophages and amoebae was less effective in eliminating the porin mutants compared to the wild-type. The cell wall permeability has been shown to influence the susceptibility of mycobacteria to host antimicrobial molecules such as defensins and lysozyme (Gao et al., 2003). Differences in the complement of porins of highly pathogenic slow-growing mycobacteria and the less-pathogenic fast-growing mycobacteria may therefore be one of the factors accounting for their divergent intracellular persistence. Similarly, the acidification of the phagosome takes place in both macrophages and amoebae (McNeil et al., 1983
). In contrast to the members of the M. tuberculosis complex, M. smegmatis cannot prevent the acidification of the phagosome. After 5 h, M. smegmatis-containing phagosomes exhibit a pH of 5·2 (Cotter & Hill, 2003
). However, the enhanced intracellular survival of the M. smegmatis mutants was not caused by a higher resistance towards low phagosomal pH, because the differences in the growth rates between the three strains in broth cultures were not dependent on the pH of the medium. Other bactericidal mechanisms of phagocytic cells are the production of reactive oxygen intermediates and reactive nitrogen intermediates, and the activity of antimicrobial peptides and lysosomal enzymes. Reactive oxygen intermediates, lysosomal enzymes and antimicrobial peptides are employed by both macrophages and amoebae for the degradation of intracellular bacteria (Brooks & Schneider, 1985
; Bruhn et al., 2003
). A reduced permeability of the cell wall of the porin mutants for small hydrophilic molecules may implicate lower accessibility of antimicrobial substances and, as a consequence, better survival of the mutants compared to the parental strain.
In addition to the diffusion of molecules directly through the porin channels, the possibility of altered diffusion rates of molecules through the mycolic acid layer must also be considered. Stephan et al. (2004a) observed an increase in resistance of the
mspA mutant to hydrophobic antibiotics. They propose that the integration of porins in the lipid layer reduces the strong interactions of the mycolic acids and thereby facilitates the diffusion of hydrophobic molecules. A reduced diffusion of harmful hydrophobic substances present in the phagolysosome of phagocytic cells through the mycolic acid layer of the
mspA and the
mspA
mspC mutants may therefore also contribute to their improved intracellular persistence.
An interesting outcome of this and our previous study was the divergent effect of the amount of porins on the intracellular persistence of M. bovis BCG and M. smegmatis. While the transfer of the porin MspA into M. bovis BCG enhanced its intracellular survival (Sharbati-Tehrani et al., 2004), M. smegmatis showed better intracellular survival after deletion of one or two porin genes. These divergent effects may be due to differences between the compositions of phagosomes containing M. bovis BCG compared with those containing M. smegmatis. The chemical composition of a phagosome depends, among other factors, on the phagocytosed bacterium. The stage of maturation of the phagosome in turn determines the degree to which the resident bacteria are exposed to acidic conditions, hydrolytic enzymes, cationic antimicrobial peptides and reactive oxygen and nitrogen compounds on the one hand, and the degree to which they have access to nutrients on the other (Schnappinger et al., 2003
; Vergne et al., 2004
). The survival of an intraphagosomal bacterium depends on the balance between these antimicrobial activities and nutrient supply, which may both be influenced by the amount of porins present in the otherwise extremely impermeable cell wall of mycobacteria. Pathogenic slow-growing mycobacteria have been shown to block phagosome maturation. The cell wall constituent lipoarabinomannan (LAM) interferes with acquisition of late endosomal constituents (Fratti et al., 2003
), and by this means the bacteria are not exposed to the antimicrobial activities of the macrophage. Another glycolipid, phosphatidylinositol mannoside (PIM), on the other hand, stimulates fusion with early endosomal compartments, thereby ensuring access to endocytosed nutrients, including iron (Vergne et al., 2004
). In this situation, the advantage of an enhanced nutrient supply brought about by additional porins may outweigh the disadvantage of reduced protection against antimicrobial activities. Opportunistic fast-growing mycobacteria such as M. smegmatis reside in phagosomes, representing a very different environment. They cannot prevent the maturation of the phagosomes (Cotter & Hill, 2003
) and are accordingly heavily attacked by lysosomal components. The reduced protection against these components caused by a higher amount of porins in the cell wall of M. smegmatis may therefore outweigh the favourable effect of porins on nutrient supply.
In addition, in contrast to M. smegmatis, the recombinant expression of mspA in M. bovis BCG derivatives was clearly decreased (Mailaender et al., 2004; Sharbati-Tehrani et al., 2004
). Using the plasmid pMN013 harbouring the transcriptional fusion of the promoter pimyc with the mspA gene (see above), Mailaender et al. (2004)
have shown a 40-fold lower density of MspA-like pores in the cell wall of the recombinant M. bovis BCG compared with M. smegmatis. Our M. bovis BCG derivative, however, harboured the integrative plasmid pSSa100, including mspA with its own promoter. RT-real-time PCR data confirmed that pSSa100 confers a clearly lower expression level than pMN013 (data not shown), which is consistent with the low amounts of recombinant MspA in our M. bovis BCG derivative (Sharbati-Tehrani et al., 2004
). Because of the ability of M. bovis BCG to persist and to replicate intracellularly, the small amount of MspA pores in the cell wall of the porin-expressing M. bovis BCG may rather represent an advantage in nutrient supply for intracellular bacilli than a disadvantage due to the diffusion of harmful compounds. As mentioned above, and in contrast to M. bovis BCG, M. smegmatis is not able to inhibit phagosome maturation. Hence, the lower density of porins in the cell wall of the porin deletion mutants of M. smegmatis might contribute to a walling-off towards the hostile phagosomal environment.
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ACKNOWLEDGEMENTS |
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Received 16 February 2005;
revised 30 March 2005;
accepted 14 April 2005.
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