Nitric oxide levels regulate macrophage commitment to ... - CiteSeerX

The FASEB Journal express article 10.1096/fj.03-1450fje. Published online May 7, 2004.

Nitric oxide levels regulate macrophage commitment to apoptosis or necrosis during pneumococcal infection Helen M. Marriott,* Farzana Ali,* Robert C. Read,* Tim J. Mitchell,† Moira K. B. Whyte,* and David H. Dockrell* *Division of Genomic Medicine, University of Sheffield School of Medicine and Biomedical Sciences, Sheffield; and †Division of Infection and Immunity, Institute of Biomedical and Life Sciences, University of Glasgow, Glasgow, United Kingdom Corresponding author: D. H. Dockrell, Division of Genomic Medicine, F-Floor, University of Sheffield Medical School, Beech Hill Road, Sheffield S10 2RX, United Kingdom. E-mail: [email protected] ABSTRACT Macrophages are resistant to constitutive apoptosis, but infectious stimuli can induce either microbial or host-mediated macrophage apoptosis. Phagocytosis and killing of opsonized pneumococci by macrophages are potent stimuli for host-mediated apoptosis, but the link between pneumococcal killing and apoptosis induction remains undefined. We now show phagocytosis of pneumococci by differentiated human monocyte-derived macrophages (MDM) results in up-regulation of inducible nitric oxide synthase (iNOS) and increased production of NO and reactive nitrogen species. NO accumulation in macrophages initiates an apoptotic program that involves NO-dependent mitochondrial membrane permeabilization, Mcl-1 downregulation, and caspase activation and results in nuclear condensation and fragmentation. An inhibitor of mitochondrial permeability transition, bongkrekic acid, decreases pneumococcalassociated macrophage apoptosis. Conversely, inhibition of NO production using iNOS inhibitors decreases bacterial killing and shifts the cell death program from apoptosis to necrosis. Pneumolysin contributes to both NO production and apoptosis induction. After initial microbial killing, NO accumulation switches the macrophage phenotype from an activated cell to a cell susceptible to apoptosis. These results illustrate important roles for NO in the integration of host defense and regulation of inflammation in human macrophages. Key words: programmed cell death • human • monocytes

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he resident professional phagocytes in tissues, macrophages (Mφ), are an early sensor of infection (1). Phagocytosis of microorganisms results in a coordinated immune response involving microbial killing, regulation of the inflammatory response, and development of immune responses (2). Differentiated Mφ are long-lived cells and demonstrate constitutive resistance to apoptosis (3), but many bacterial infections are characterized by induction of Mφ apoptosis (4). In many cases, Mφ apoptosis enhances microbial pathogenesis by allowing immune evasion (5). Evidence of host-mediated Mφ apoptosis has been less apparent although chronic intracellular pathogens, most notably Mycobacterium tuberculosis, have been associated with decreased microbial replication in the presence of Mφ apoptosis (6). In these cases, the link between mycobacterial killing and apoptosis has been difficult to establish due to Page 1 of 26 (page number not for citation purposes)

the persistence of intracellular bacteria. A number of potential mediators of mycobacterialassociated Mφ apoptosis have been proposed including tumor necrosis factor-α (TNF-α; ref 7), Fas ligand (FasL; ref 8), nitric oxide (NO; ref 9), and activation of the P2X7 purinergic receptor (6), but discordant results have been reported with regard to whether Mφ apoptosis induced by each factor also limits mycobacterial replication. Streptococcus pneumoniae, the pneumococcus, is a significant human pathogen and a common cause of community-acquired pneumonia, meningitis, and otitis media (10). Mφ phagocytose and kill opsonized pneumococci (11). Streptococci, including the group B streptococcus and pneumococci, induce Mφ apoptosis (12, 13). We have demonstrated that an unusual feature of pneumococcal-associated Mφ apoptosis, in comparison to other examples of Mφ apoptosis during acute bacterial infection, is its relationship to the intracellular burden of opsonized pneumococci and intracellular killing (13, 14). A role for Mφ apoptosis in host defense has been suggested in vitro and in vivo by the observation that inhibition of caspase activation results in decreased pneumococcal killing as well as decreased Mφ apoptosis (14, 15). NO is a potential mediator of both antimicrobial host defense (16) and apoptosis induction during exposure of Mφ to pneumococci. During infection, rodent Mφ express inducible NO synthase (iNOS/NOS2) and produce NO (16, 17). iNOS or NO has also been demonstrated in human monocytes/Mφ (18–20). NO contributes to the initial microbicidal activity in vitro and in vivo against intracellular pathogens including Salmonella typhimurium (21, 22), M. tuberculosis (23), and Lesihmania chagasi (24). Pneumococci have been demonstrated to induce NO production in murine Mφ in vitro (25, 26). NO also plays an important role in the regulation of cell survival and can both enhance (27, 28) and inhibit apoptosis (29), depending on the level of NO, cell type, and duration of exposure. We have, therefore, investigated the role of NO in pneumococcal-associated Mφ apoptosis and demonstrate that NO production in response to pneumolysin induces mitochondrial membrane permeabilization and apoptosis but that iNOS inhibition induces cell necrosis with decreased bacterial killing. MATERIALS AND METHODS Construction of pneumolysin negative mutant Plasmid pLC1 contains the pneumolysin gene in PCR-Script vector (Invitrogen). This plasmid was used for inverse PCR using primers INV1 (AGGCGCGCCTTACAGCTTACAGAAACGGAGATT) and INV2 (GACGGCGCGCCTGATGCTACTTATCCAAC). Sites for the restriction enzyme AscI were introduced on the primers (underlined). After inverse PCR, the product (containing the 5′ and 3′ regions of the pneumolysin gene and the pPCR-Script vector) was purified, restricted with AscI, self-ligated, and recombinant plasmid isolated. This plasmid was linearized with ASCII, and an erythromycin resistance cassette with AscI ends was ligated into the construct. The resulting plasmid (pLC2) contains the erythromycin resistance gene flanked on either side by ~300 bp of 5′ and 3′ sequence from the pneumolysin gene. pLC2 was used to transform Streptococcus pneumoniae D39. Gene disruptions were selected by plating the transformation mix on 1 µg erythromycin. PCR and DNA sequencing confirmed disruption of the pneumolysin gene. Bacteria Type 1 (WHO reference laboratory strain SSISP; Statens Seruminstitut), type 2 (D39 strain, Page 2 of 26 (page number not for citation purposes)

NCTC7466), or pneumolysin deficient S. pneumoniae (pneumococci) were grown as described (11), thawed aliquots were opsonized, and bacterial numbers were confirmed by the surface viable count method after inoculation on blood agar (13). Purified pneumolysin was prepared as described previously (25). Cells and infection Peripheral blood mononuclear cells were isolated by density centrifugation from whole blood donated by healthy volunteers as described previously (13); 14 day monocyte-derived macrophages (MDM) were infected with opsonized pneumococci at a multiplicity of infection of 10 or mock infected (13). In certain experiments, 50 µM 1400W (Calbiochem) as a specific iNOS inhibitor (30), 500 µM L-NG-methyl arginine (L-NMA, Sigma) as a general NOS inhibitor, NOC-18 (Alexis) as a NO donor, bongrekic acid as an inhibitor of mitochondrial permeability transition at 5-50 µM (Sigma) (31), or purified pneumolysin (25) was added 1 h before infection and at 4 h. In experiments where exogenous pneumolysin was added alone, addition was at 0 h. SDS-PAGE and Western immunoblotting Whole cell extracts were lysed in buffer (20 mM Tris-HCl (pH 7.4), 150 mM NaCl, 5 mM EDTA, 5 mM EGTA, 1% SDS, 1:25 protease inhibitor (Boehringer-Mannheim). Cytosolic fractions were extracted exactly as described (32), and a lack of mitochondrial contamination was confirmed by absence of Cox IV detection (clone 20E8, mouse monoclonal, Molecular Probes) despite detection of a band of appropriate size in mitochondrial fractions. Protein was quantified using a modified Lowry protocol (DC protein assay, Bio-Rad), and equal protein was loaded per lane. Samples were separated by SDS-PAGE (8-15%) and blotted onto PVDF membranes (Millipore) for detection of iNOS or Trans-blot nitrocellulose membranes (Bio-Rad) for detection of cytochrome c and Mcl-1. Ponceau S staining confirmed protein transfer and equal loading, as did incubation with antibody against actin (goat polyclonal, 1:500, Santa Cruz Biotechnology or rabbit polyclonal. 1:2000, Sigma). Blots were incubated at 4°C for 16 h with antibodies to iNOS (clone 54, mouse monoclonal, 1:2,000, BD Biosciences), cytochrome c (clone 7H8.2C12, mouse monoclonal, 1:1000, BD Biosciences), or Mcl-1 (S-19, rabbit polyclonal, 1:500 Santa Cruz Biotechnology). Protein detection was with horseradish peroxidase conjugated secondary antibodies (Dako) and ECL (Amersham Pharmacia). NO detection NO production was assessed by three techniques, indirectly by extracellular nitrite and intracellular nitrotyrosine, and directly by the increase of fluorescence of the intracellular dye 4amino-5-methylamino-2′,7′-difluorescein diacetate (DAF-FM, Molecular Probes). Nitrite Extracellular nitrite was measured using triplicate samples from MDM culture supernatants. Nitrite was quantified by O3/chemiluminescence (Model Mk2B; Glaxo-Smith Kline) using reduction of nitrite to NO and comparison to known nitrite standards, as previously reported (33).

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DAF-FM DAF-FM fluorescence was measured by FACS® analysis; 1 x 106 cells were incubated in RPMI without phenol red (Invitrogen) for 30 min at 37°C and incubated with 5 µM DAF-FM for 30 min (34). Cells were then incubated in RPMI minus phenol red for 30 min at 37°C, washed, and scraped. Cells were analyzed on a FACSCalibur® flow cytometer (BD Biosciences); 30,000 events were saved and analyzed using Cell Quest software® (BD Biosciences). Change in fluorescence intensity (∆MFI) was calculated by subtracting unstained from stained cells. To exclude any effect of extracellular Ca2+ or Mg2+ on fluorescence signal, controls were performed with DAF-FM incubation in the presence of 5 mM EDTA (35). MDM grown on coverslips stained with DAF-FM as above were also imaged by confocal (Leica TCS SP2 system) or fluorescence microscopy (Nikon Diaphot 300). As a positive control, MDM were incubated with 100 µg/ml of NOC-18 for 1 h. In costaining experiments, cells were incubated in 500 nM MitoTracker Red 580 (Molecular Probes) or 75 nM LysoTracker Red DND-99 (Molecular Probes) for 30 min at 37°C before incubation with DAF-FM or with 2.5 µg/ml of propidium iodide (Sigma) in H2O for 15 min at 20°C after incubation with DAF-FM. Cells were imaged by fluorescence microscopy (Nikon Diaphot 300). Nitrotyrosine To detect nitrotyrosine MDM grown on coverslips were washed, fixed with ethanol:acetic acid (95:5) for 1 min, washed, and incubated for 1 h at 20°C with 1% BSA (First Link, UK Ltd) in PBS. After being washed, MDM were incubated with 10 µg/ml anti-nitrotyrosine (Upstate) in 1% BSA in PBS for 2 h at 20°C, washed, and incubated with an anti-mouse FITC conjugated secondary antibody (1:100, Sigma) for 75 min at 20°C. Coverslips were washed and mounted on glass slides, with Vectashield (Vector Laboratories), and cells were imaged by confocal microscopy. Bacterial killing assays The 6 h intracellular killing assay was performed as described previously (13). For the 16 h intracellular killing assay, MDM were washed at 4 h postinfection, incubated with 40 Mu benzylpenicillin and 20 µg/ml gentamycin for 30 min at 37°C, washed, and incubated with 0.75 µg/ml vancomycin (Sigma), until lysed, as for the 6 h assay. Quadruplicate wells were lysed for each data point. Apoptosis detection Nuclear features of MDM apoptosis detected by 4’6’-diamidino-2-phenylindole (DAPI, Molecular Probes), TUNEL staining (Intergen) with propidium iodide counter-staining and fluorescence microscopy to detect bacterial internalization were performed as previously reported (13). To detect loss of ∆ψm, MDM were stained in 10 µM 5,5′,6,6′-tetrachloro-1,1′,3,3′ tetraethylbenzimidazolocarbocyanine iodide (JC-1; Molecular Probes) for 30 min at 37°C. Coverslips were mounted and examined by fluorescent microscopy. Loss of ∆ψm was confirmed by loss of red punctate staining. In these experiments, MDM pretreated with 5 µM staurosporine (Sigma) for 2 h were used as a positive control for loss of ∆ψm. Caspase activation was determined by flow cytometry after staining with FITC-VAD-FMK (Promega). MDM were incubated with 10 µM FITC-VAD-FMK for 20 min at 37°C, followed by staining for necrotic Page 4 of 26 (page number not for citation purposes)

cells with 2.5 µg/ml propidium iodide at room temperature for 15 min. Cell density was determined by counting at least 300 cells in duplicate samples after DAPI staining and calculating the mean number of cells per field when examined with x100 objective. Quadrant position for flow cytometric assessment of necrosis was checked using unstained cells as negative controls and cells permeabilized with 1% Saponin as positive controls. Statistics All results are recorded as means ± SE unless otherwise stated. Nonparametric testing was performed as described using Prism 3.03 software (GraphPad Inc.). Significance was defined as P < 0.05. RESULTS Infection with pneumococci increases NO production by MDM Phagocytosis of pneumococci by human MDM resulted in increased production of both NO and nitrotyrosine (a reactive nitrogen species; RNS; Fig. 1A). Confocal microscopy revealed a punctate intracellular NO staining pattern in MDM exposed to pneumococci. NO production in MDM was further demonstrated by flow cytometry using DAF-FM (Fig. 1B). These results were reproduced for both type 1 and type 2 pneumococci (see below). Increased NO production in infected as compared with mock infected MDM was detectable 6 h after infection, and NO production was further increased by 12 h after infection (Fig. 1B, C). Nitrite levels 8 h after infection in pneumococcal-infected as opposed to mock-infected MDM were 1.43 ± 0.51 vs. 0.67 ± 0.20 µM, n = 6, P < 0.05 (means±SE). Western blotting on cytosolic extracts revealed that even though iNOS was detectable in mock-infected cultures, suggesting activation of control Mφ by in vitro culture, iNOS protein was initially up-regulated in MDM after pneumococcal infection (compare iNOS protein expression at 12 h after pneumococcal infection to that at 4 h or to levels after mock infection; Fig. 1D) but was subsequently down-regulated (iNOS expression at 20 h after pneumococcal infection (Fig. 1D). By 20 h postinfection, however, iNOS was down-regulated in pneumococcal-infected MDM. In addition, iNOS inhibition significantly inhibited production of nitrite, an extracellular marker of NO production, in culture supernatants (Fig. 1E). Intracellular localization of NO in MDM after phagocytes of pneumococci To determine whether NO production was localized to a specific organelle, dual staining for specific organelles and for NO was performed (Fig. 2). Since pneumococci traffic to phagolysosomes (11), we investigated whether NO localized to lysosomes, particularly as these organelles have been proposed as a site of NO production (36). Staining of lysosomes was most prominent up to 16 h and then decreased, but intensity of NO staining increased steadily up to 20 h in many donors. Costaining experiments were performed at 14 h and showed NO staining in areas adjacent to but typically distinct from lysosomes (Fig. 2A). At this time point, DAF-FM staining was more diffuse than at later time points with only small areas of punctate staining. Similar results were obtained when we studied the localization of NO relative to areas containing pneumococci (Fig. 2B). Once again, NO and bacteria were adjacent to each other but distinct. The images shown were obtained at 16 h after infection and demonstrate that NO staining was greater at this later time point, with larger punctate areas apparent in the cytoplasm. Page 5 of 26 (page number not for citation purposes)

Mitochondria have been suggested as a potential source of NO production under specific circumstances (37), but mitochondrial staining also did not colocalize with NO (Fig. 2C). NO contributes to killing of pneumococci in MDM To examine the contribution of NO to the anti-microbial activity of MDM against pneumococci, bacterial viability was assessed at 6 and 16 h postinfection. After internalization by macrophages, pneumococci do not replicate and bacterial viability is slowly reduced (11). As shown in Fig. 3A, iNOS inhibition did not alter the initial numbers of internalized bacteria 4 h after infection. However, although there was a steady fall in bacterial viability with time for all cultures (Fig. 3B), iNOS inhibition slowed the decline in viability of intracellular bacteria, illustrating that NO contributes to the decline in bacterial viability at both time points. Furthermore, the majority (75%) of cultures treated with vehicle control had no viable intracellular bacteria at 16 h, whereas the majority (63%) of those treated with the iNOS inhibitor had viable intracellular bacteria. Addition of an exogenous NO donor, NOC-18, significantly decreased intracellular bacterial viability 6 h after infection by a mean of 62% (n=7) but at concentrations ranging between 1-100 µg/ml had no impact on the number of internalized bacteria inside MDM 4 h after infection: means ± SE bacteria/MDM 1.7 ± 0.3- 1.8 ± 0.4 as compared with 1.7 ± 0.4 for mock-infected cultures (n=6). NO production contributes to pneumococcal-associated MDM apoptosis Internalization and killing of pneumococci are associated with apoptosis induction (13). Apoptosis of directly infected MDM is Fas-independent (13) and is not enhanced by exogenous TNF-α (our unpublished observation). Since NO has the ability to contribute to the regulation of cell lifespan in other settings, we hypothesized that NO production could provide a link between pneumococcal killing and MDM apoptosis. As shown in Fig. 4A and B, the number of TUNEL+ cells after pneumococcal infection was significantly decreased by iNOS inhibition. Similar results were obtained when nuclear features of apoptosis were quantified after treatment with two different iNOS inhibitors (Fig. 4C). However, NO alone was not sufficient to explain apoptosis induction in MDM since an exogenous NO donor, NOC-18, did not significantly increase MDM apoptosis in mock-infected cells and only caused a significant increase in apoptosis in MDM cultures infected with pneumococci, Fig. 4D. The increase in median MDM apoptosis observed with the combination of infection and 100 µg/ml NOC-18 (+21%) was greater than observed for either 100 µg/ml NOC-18 (+5%) or pneumococcal infection alone (+12%), suggesting that the combination of pneumococci and NO are required for optimal apoptosis induction. NO production during pneumococcal infection determines the mechanism of MDM cell death To further explore how NO contributes to engagement of an apoptotic program in MDM incubated with pneumococci, we examined the mechanism of cell death. Apoptosis may be mediated by death receptors or can be initiated by mitochondrial membrane permeabilization (MMP; ref 38). Pneumococcal-associated MDM apoptosis is associated with two features of MMP. Loss of mitochondrial inner transmembrane potential (∆ψm; Fig. 5A and B) was observed for all donors exposed to pneumococci and was abrogated by iNOS inhibition. We quantified the percentage of cells with loss of ∆ψm for a series of donors and were able to demonstrate that, in Page 6 of 26 (page number not for citation purposes)

addition to decreasing the percentage of MDM with nuclear features of apoptosis after pneumococcal infection, iNOS inhibition also resulted in a significant decrease in the number of cells demonstrating loss of ∆ψm (Fig. 5C). Pneumococcal-associated MDM apoptosis is also characterized by cytosolic translocation of cytochrome c (Fig. 5D), and inhibition of iNOS decreased cytochrome c translocation. Mφ survival has been documented to be critically dependent on expression of the anti-apoptotic Bcl-2 family member myeloid cell leukaemia sequence (Mcl)-1 (3). Mcl-1 down-regulation during pneumococcal-associated MDM apoptosis is abrogated by iNOS inhibition (Fig. 5E). To determine whether MMP was essential for pneumococcal-associated MDM apoptosis, we treated cultures with bongkrekic acid. Bongkrekic acid binds to adenine nucleotide translocator (ANT) and inhibits mitochondrial permeability transition induced by pro-apoptotic Bcl-2 family members such as Bax and Bid resulting in decreased MMP and apoptosis (31, 39). As shown in Fig. 6, bongkrekic acid decreased pneumococcal-associated macrophage apoptosis, suggesting MMP is a critical feature of pneumococcal-associated MDM apoptosis. Inhibition of iNOS does not prevent MDM cell death The decreased MMP and apoptosis observed after iNOS inhibition of pneumococcal-infected MDM cultures could be the result of increased overall cell survival or could reflect an alternative pathway of cell death that is not associated with the late features of apoptosis such as DNA cleavage and nuclear fragmentation. Despite evidence of decreased MMP and of maintenance of Mcl-1 protein levels, iNOS inhibition did not prevent cell death. Cell density decreased significantly despite iNOS inhibition in pneumococcal-infected cultures, just as was observed in pneumococcal-infected cultures in the absence of iNOS inhibition (Fig. 7A). To exclude the possibility that differences in cell number reflected differing kinetics of apoptosis, we examined levels of apoptosis by DAPI staining at various times from 8-48 h after pneumococcal infection and confirmed that there was no evidence of an earlier onset of apoptosis in pneumococcalinfected MDM in the presence of iNOS inhibition as compared with cultures without iNOS inhibition (data not shown). Cells that demonstrate necrosis lose membrane integrity, in contrast to apoptotic cells that have intact membrane integrity until relatively late stages of the apoptotic process. In the presence of iNOS inhibition, pneumococcal-infected MDM showed loss of membrane integrity (as determined by inability to exclude high levels of propidium iodide), as compared with infected cultures in the absence of iNOS inhibition (which were able to exclude high levels of propidium iodide), consistent with increased cell death by necrosis in the presence of iNOS inhibition (Fig. 7B). In addition, iNOS inhibition prevented the caspase activation observed during pneumococcal-associated MDM apoptosis. These results illustrate the important role of NO in regulating the mechanisms and type of cell death after exposure to pneumococci. Role of pneumolysin in pneumococcal-associated MDM apoptosis Since pneumolysin has been previously demonstrated to contribute to neuronal apoptosis during meningitis (40) and has been shown to stimulate NO production by murine macrophages (25), we investigated its role in the current model. Pneumolysin can induce MDM apoptosis (Fig. 8A) but does not result in comparable levels of apoptosis to that observed with internalization of whole bacteria (Fig. 8B). However, internalization of pneumolysin-deficient pneumococci results in significantly less MDM apoptosis than internalization of the parental strain (Fig. 8B). Under these conditions, infection of MDM with the pneumolysin deficient strain and exogenous Page 7 of 26 (page number not for citation purposes)

pneumolysin results in a comparable level of MDM apoptosis to infection with the parental strain. The pneumolysin deficient strain also resulted in decreased MMP (Fig. 8C). Pneumolysin also played a role in NO production by MDM, as the pneumolysin- deficient strain induced less NO production than its parental strain (Fig. 8D). DISCUSSION NO production in human Mφ exposed to pneumococci We have demonstrated that after infection of human Mφ with pneumococci, iNOS is upregulated and there is enhanced production of NO and RNS. NO was detected using DAF-FM as a marker of intracellular NO production (41). Specificity for NO detection was confirmed by documenting an inhibition of signal with selective iNOS inhibition (42, 43). Nitrite is a commonly used surrogate marker for NO production in human cells, and we measured nitrite by O3/chemiluminescence, a highly sensitive technique. In addition, we also detected evidence of up-regulation of iNOS protein and increased nitrotyrosine production in pneumococcal-infected MDM. The level of nitrite we have detected is less than that generally induced in rodent Mφ (44) and supports the concept that the levels of NO produced by human Mφ are lower (24). In addition, we found evidence for spatial regulation of NO, with a punctate cytoplasmic localization similar to that described in murine Mφ, where the pattern corresponds to localization of iNOS in 50-80 nm microvesicles distinct from other cellular organelles (45, 46). NO contributes to loss of viability of intracellular pneumococci within the first 6 h of infection. Even the low levels of NO produced play a significant role in loss of bacterial viability, although other factors are likely to contribute since iNOS inhibition only partially blocks loss of viability and intracellular colony counts continue to fall after iNOS inhibition. Antimicrobial properties of NO include peroxidation of bacterial membrane lipids, inactivation of critical enzymes, inhibition of protein synthesis, induction of DNA mutation, and inhibition of DNA repair (16, 47). These activities are, however, enhanced by many other molecules including the interaction with reactive oxygen species (ROS; ref 48). For example, in murine Mφ NO synergizes with ROS in the early killing of S. typhimurium (21, 22). These anti-microbial molecules must be spatially and temporally regulated to prevent cellular toxicity both to the directly infected Mφ and to bystander cells. NO-mediated Mφ apoptosis during pneumococcal infection Apoptosis of MDM after in vitro exposure to pneumococci is a delayed response and does not occur during the initial period of intracellular killing (14). At later time points after infection, we observed that NO played a critical role in determining the fate of Mφ. NO induced MMPdependent apoptosis, in association with down-regulation of a critical regulator of Mφ survival, Mcl-1 (3). MMP was demonstrated to be a central feature of pneumococcal-associated Mφ apoptosis rather than part of an amplification loop since an inhibitor of mitochondrial permeability transition and MMP reduced apoptosis. NO has been demonstrated to both inhibit and induce apoptosis, and the variability in results reflects differences in levels of NO, cell type, and activation state, but recent studies have clarified these findings (49). Sustained low level production of NO inhibits Mφ apoptosis whereas acute high-level production induces apoptosis (49). Page 8 of 26 (page number not for citation purposes)

An explanation for the biphasic effect of NO on cell survival has been provided in Jurkat T cells (28). During accumulating nitrosative stress, NO initially preserves ∆ψm. Greater levels of nitrosative stress in association with oxidative stress cause loss of ∆ψm and therefore MMP (50). Our study extends previous observations by studying the effect of NO on Mφ apoptosis in a physiologic situation. Our model using highly differentiated human Mφ internalizing and killing opsonized pneumococci suggests apoptosis is absent during initial NO accumulation but is induced after sustained NO accumulation. In this setting, however, loss of ∆ψm is a feature of NO-mediated apoptosis, in contrast to the increase in ∆ψm observed in Mφ treated with high concentrations of NO donors in the absence of infection (51). These differences may reflect the effects of altered signal transduction pathways associated with pneumococcal infection or differences in the cell types used in the studies. Pneumolysin was necessary but not sufficient for optimal Mφ apoptosis induction that required internalization of whole bacteria. As was the case for exogenous NO, additional signals produced during the process of phagocytosis and killing of bacteria were necessary for maximal Mφ apoptosis. This contrasts with neuronal cells, in which apoptosis induction was mediated by extracellular pneumolysin, and argues for the existence of a separate mechanism of apoptosis induction in Mφ exposed to pneumococci (40). It also suggests that during the phagocytosis of live bacteria multiple competing signals may combine to initiate an apoptotic program. Shifting the input of one of these contributing factors, as we have demonstrated in this study by decreasing NO production, can have a major effect on the type of cell death observed. Inhibition of iNOS during pneumococcal infection decreases MMP and Mcl-1 down-regulation but induces a necrotic cell death. Blocking NO production by iNOS had varying effects on different parameters measured. There was greater inhibition of nuclear apoptosis and cytochrome c translocation than there was inhibition of loss of ∆ψm. This may in part be explicable by the super-imposed effects of cell necrosis on mitochondria (38). Induction of necrosis in the presence of iNOS inhibition is likely the result of ATP depletion in association with a decrease in cytosolic pH, a feature of phagocytes that contain large numbers of intracellular bacteria (52). The level of NO therefore has an important role in determining the mechanism of cell death during pneumococcal infection of Mφ. Potential significance of host-mediated Mφ apoptosis induced by NO NO can, under certain circumstances, decrease the development of inflammation in the lung and can decrease chemotaxis by modulating the response to chemokines and the expression of integrins (53–55). Induction of Mφ apoptosis represents a further example of how NO may regulate resolution of inflammation after infection, by removing activated cells and limiting tissue damage induced by production of pro-inflammatory factors, such as the cytokines produced by activated Mφ after phagocytosis of bacteria (2) and high levels of NO (56). In addition to aiding resolution of inflammation, Mφ apoptosis provides a source of antigens for presentation by dendritic cells, hence linking innate and adaptive immunity (57). Host-mediated Mφ apoptosis may have a particular role in microbial killing. The importance of Mφ apoptosis in controlling microbial replication is best established for mycobacteria (58). In this case, apoptosis but not necrosis can help limit intracellular replication. Mycobacteria upregulate anti-apoptotic molecules including Mcl-1 to prevent host-mediated Mφ apoptosis (59). Page 9 of 26 (page number not for citation purposes)

Although NO has been implicated in mycobacterial-associated Mφ apoptosis previously (9), its role has been difficult to establish due to the chronic nature of mycobacterial infection of Mφ and the inability of other groups to confirm the role of NO (6). Nevertheless, the importance of NO in host defense against mycobacteria is well established as supported by the number of Mφ genes whose expression is modified by NO during infection with M. tuberculosis (60) and the existence of specific genes in M. tuberculosis which protect against nitrosative stress (61). In contrast, pneumococcal infection of Mφ is acute and phagocytosed bacteria are completely cleared (13). Pneumococci have not been demonstrated to possess genes conferring resistance to nitrosative stress, and therefore pneumococcal infection of Mφ may be a more effective example of NO-induced host-mediated Mφ apoptosis. Previous studies of murine pneumonia have failed to document an association between pneumococcal killing and levels of NO (62, 63). However, those studies focused on established infection, with marked inflammatory damage to the lung, and under these circumstances the Mφ host defense has been overwhelmed by high numbers of bacteria (15). We have documented that during subclinical pneumococcal infection Mφ apoptosis contributes to host defense (15), and dysregulated NO, ROS, and pro-inflammatory cytokine production is likely to worsen outcome under these circumstances. A beneficial role for Mφ-produced NO is therefore most likely during exposure to small numbers of bacteria as might occur during subclinical infection. CONCLUSIONS We provide evidence of the expanding role of NO in host defense by using the physiological model of Mφ phagocytosing and killing pneumococci. At lower concentrations, NO contributes to pneumococcal killing, but at higher concentrations, it facilitates the transition from an activated, apoptosis-resistant Mφ phenotype into a phenotype susceptible to apoptosis. Inhibition of iNOS shifts the cell death program from MMP-mediated apoptosis to necrosis. These findings implicate NO as an important factor in host-mediated Mφ apoptosis during pneumococcal infection and suggest its presence allows removal of activated Mφ by an apoptotic process that is likely to minimize local inflammatory damage to tissue. ACKNOWLEDGMENTS This work was supported by a Wellcome Trust Advanced Clinical Fellowship to D. H. Dockrell (#065054). The authors are grateful to Patrick Vallance and Ian Sabroe for critical comments during the preparation of this manuscript and to Ian Palmer for assistance with cell imaging. REFERENCES 1.

Sibille, Y., and Reynolds, H. Y. (1990) Macrophages and polymorphonuclear neutrophils in lung defense and injury. Am. Rev. Respir. Dis. 141, 471–501

2.

Underhill, D. M., and Ozinsky, A. (2002) Phagocytosis of microbes: complexity in action. Annu. Rev. Immunol. 20, 825–852

3.

Liu, H., Perlman, H., Pagliari, L. J., and Pope, R. M. (2001) Constitutively activated Akt-1 is vital for the survival of human monocyte-differentiated macrophages. Role of Mcl-1, independent of nuclear factor (NF)-kappaB, Bad, or caspase activation. J. Exp. Med. 194, Page 10 of 26 (page number not for citation purposes)

113–126 4.

Zychlinsky, A., and Sansonetti, P. (1997) Perspectives series: host/pathogen interactions. Apoptosis in bacterial pathogenesis. J. Clin. Invest. 100, 493–495

5.

Gao, L. Y., and Kwaik, Y. A. (2000) The modulation of host cell apoptosis by intracellular bacterial pathogens. Trends Microbiol. 8, 306–313

6.

Lammas, D. A., Stober, C., Harvey, C. J., Kendrick, N., Panchalingam, S., and Kumararatne, D. S. (1997) ATP-induced killing of mycobacteria by human macrophages is mediated by purinergic P2Z(P2X7) receptors. Immunity 7, 433–444

7.

Keane, J., Balcewicz-Sablinska, M. K., Remold, H. G., Chupp, G. L., Meek, B. B., Fenton, M. J., and Kornfeld, H. (1997) Infection by Mycobacterium tuberculosis promotes human alveolar macrophage apoptosis. Infect. Immun. 65, 298–304

8.

Oddo, M., Renno, T., Attinger, A., Bakker, T., MacDonald, H. R., and Meylan, P. R. (1998) Fas ligand-induced apoptosis of infected human macrophages reduces the viability of intracellular Mycobacterium tuberculosis. J. Immunol. 160, 5448–5454

9.

Rojas, M., Barrera, L. F., Puzo, G., and Garcia, L. F. (1997) Differential induction of apoptosis by virulent Mycobacterium tuberculosis in resistant and susceptible murine macrophages: role of nitric oxide and mycobacterial products. J. Immunol. 159, 1352–1361

10. Tuomanen, E. I., Austrian, R., and Masure, H. R. (1995) Pathogenesis of pneumococcal infection. N. Engl. J. Med. 332, 1280–1284 11. Gordon, S. B., Irving, G. R., Lawson, R. A., Lee, M. E., and Read, R. C. (2000) Intracellular trafficking and killing of Streptococcus pneumoniae by human alveolar macrophages are influenced by opsonins. Infect. Immun. 68, 2286–2293 12. Ulett, G. C., Bohnsack, J. F., Armstrong, J., and Adderson, E. E. (2003) Beta-hemolysinindependent induction of apoptosis of macrophages infected with serotype III group B streptococcus. J. Infect. Dis. 188, 1049–1053 13. Dockrell, D. H., Lee, M., Lynch, D. H., and Read, R. C. (2001) Immune-mediated phagocytosis and killing of Streptococcus pneumoniae are associated with direct and bystander macrophage apoptosis. J. Infect. Dis. 184, 713–722 14. Ali, F., Lee, M. E., Iannelli, F., Pozzi, G., Mitchell, T. J., Read, R. C., and Dockrell, D. H. (2003) Streptococcus pneumoniae-associated human macrophage apoptosis after bacterial internalization via complement and Fcgamma receptors correlates with intracellular bacterial load. J. Infect. Dis. 188, 1119–1131 15. Dockrell, D. H., Marriott, H. M., Prince, L. R., Ridger, V. C., Ince, P. G., Hellewell, P. G., and Whyte, M. K. (2003) Alveolar macrophage apoptosis contributes to pneumococcal clearance in a resolving model of pulmonary infection. J. Immunol. 171, 5380–5388 16. Bogdan, C. (2001) Nitric oxide and the immune response. Nat. Immunol. 2, 907–916 Page 11 of 26 (page number not for citation purposes)

17. MacMicking, J., Xie, Q. W., and Nathan, C. (1997) Nitric oxide and macrophage function. Annu. Rev. Immunol. 15, 323–350 18. Reiling, N., Ulmer, A. J., Duchrow, M., Ernst, M., Flad, H. D., and Hauschildt, S. (1994) Nitric oxide synthase: mRNA expression of different isoforms in human monocytes/macrophages. Eur. J. Immunol. 24, 1941–1944 19. Dumarey, C. H., Labrousse, V., Rastogi, N., Vargaftig, B. B., and Bachelet, M. (1994) Selective Mycobacterium avium-induced production of nitric oxide by human monocytederived macrophages. J. Leukoc. Biol. 56, 36–40 20. Nicholson, S., Bonecini-Almeida Mda, G., Lapa e Silva, J. R., Nathan, C., Xie, Q. W., Mumford, R., Weidner, J. R., Calaycay, J., Geng, J., Boechat, N., et al. (1996) Inducible nitric oxide synthase in pulmonary alveolar macrophages from patients with tuberculosis. J. Exp. Med. 183, 2293–2302 21. Vazquez-Torres, A., Jones-Carson, J., Mastroeni, P., Ischiropoulos, H., and Fang, F. C. (2000) Antimicrobial actions of the NADPH phagocyte oxidase and inducible nitric oxide synthase in experimental salmonellosis. I. Effects on microbial killing by activated peritoneal macrophages in vitro. J. Exp. Med. 192, 227–236 22. Mastroeni, P., Vazquez-Torres, A., Fang, F. C., Xu, Y., Khan, S., Hormaeche, C. E., and Dougan, G. (2000) Antimicrobial actions of the NADPH phagocyte oxidase and inducible nitric oxide synthase in experimental salmonellosis. II. Effects on microbial proliferation and host survival in vivo. J. Exp. Med. 192, 237–248 23. MacMicking, J. D., North, R. J., LaCourse, R., Mudgett, J. S., Shah, S. K., and Nathan, C. F. (1997) Identification of nitric oxide synthase as a protective locus against tuberculosis. Proc. Natl. Acad. Sci. USA 94, 5243–5248 24. Gantt, K. R., Goldman, T. L., McCormick, M. L., Miller, M. A., Jeronimo, S. M., Nascimento, E. T., Britigan, B. E., and Wilson, M. E. (2001) Oxidative responses of human and murine macrophages during phagocytosis of Leishmania chagasi. J. Immunol. 167, 893– 901 25. Braun, J. S., Novak, R., Gao, G., Murray, P. J., and Shenep, J. L. (1999) Pneumolysin, a protein toxin of Streptococcus pneumoniae, induces nitric oxide production from macrophages. Infect. Immun. 67, 3750–3756 26. Orman, K. L., Shenep, J. L., and English, B. K. (1998) Pneumococci stimulate the production of the inducible nitric oxide synthase and nitric oxide by murine macrophages. J. Infect. Dis. 178, 1649–1657 27. Brune, B., von Knethen, A., and Sandau, K. B. (1999) Nitric oxide (NO): an effector of apoptosis. Cell Death Differ. 6, 969–975 28. Beltran, B., Mathur, A., Duchen, M. R., Erusalimsky, J. D., and Moncada, S. (2000) The effect of nitric oxide on cell respiration: A key to understanding its role in cell survival or death. Proc. Natl. Acad. Sci. USA 97, 14602–14607 Page 12 of 26 (page number not for citation purposes)

29. Liu, L., and Stamler, J. S. (1999) NO: an inhibitor of cell death. Cell Death Differ. 6, 937– 942 30. Garvey, E. P., Oplinger, J. A., Furfine, E. S., Kiff, R. J., Laszlo, F., Whittle, B. J., and Knowles, R. G. (1997) 1400W is a slow, tight binding, and highly selective inhibitor of inducible nitric-oxide synthase in vitro and in vivo. J. Biol. Chem. 272, 4959–4963 31. Marzo, I., Brenner, C., Zamzami, N., Jurgensmeier, J. M., Susin, S. A., Vieira, H. L., Prevost, M. C., Xie, Z., Matsuyama, S., Reed, J. C., et al. (1998) Bax and adenine nucleotide translocator cooperate in the mitochondrial control of apoptosis. Science 281, 2027–2031 32. Heinloth, A., Brune, B., Fischer, B., and Galle, J. (2002) Nitric oxide prevents oxidised LDL-induced p53 accumulation, cytochrome c translocation, and apoptosis in macrophages via guanylate cyclase stimulation. Atherosclerosis 162, 93–101 33. Stevanin, T. M., Poole, R. K., Demoncheaux, E. A., and Read, R. C. (2002) Flavohemoglobin Hmp protects Salmonella enterica serovar typhimurium from nitric oxiderelated killing by human macrophages. Infect. Immun. 70, 4399–4405 34. Kojima, H., Nakatsubo, N., Kikuchi, K., Kawahara, S., Kirino, Y., Nagoshi, H., Hirata, Y., and Nagano, T. (1998) Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Anal. Chem. 70, 2446–2453 35. Broillet, M., Randin, O., and Chatton, J. (2001) Photoactivation and calcium sensitivity of the fluorescent NO indicator 4,5-diaminofluorescein (DAF-2): implications for cellular NO imaging. FEBS Lett. 491, 227–232 36. Nathan, C. (1995) Natural resistance and nitric oxide. Cell 82, 873–876 37. Zanella, B., Calonghi, N., Pagnotta, E., Masotti, L., and Guarnieri, C. (2002) Mitochondrial nitric oxide localization in H9c2 cells revealed by confocal microscopy. Biochem. Biophys. Res. Commun. 290, 1010–1014 38. Green, D. R., and Reed, J. C. (1998) Mitochondria and apoptosis. Science 281, 1309–1312 39. Zamzami, N., El Hamel, C., Maisse, C., Brenner, C., Munoz-Pinedo, C., Belzacq, A. S., Costantini, P., Vieira, H., Loeffler, M., Molle, G., et al. (2000) Bid acts on the permeability transition pore complex to induce apoptosis. Oncogene 19, 6342–6350 40. Braun, J. S., Sublett, J. E., Freyer, D., Mitchell, T. J., Cleveland, J. L., Tuomanen, E. I., and Weber, J. R. (2002) Pneumococcal pneumolysin and H(2)O(2) mediate brain cell apoptosis during meningitis. J. Clin. Invest. 109, 19–27 41. Lacza, Z., Snipes, J. A., Zhang, J., Horvath, E. M., Figueroa, J. P., Szabo, C., and Busija, D. W. (2003) Mitochondrial nitric oxide synthase is not eNOS, nNOS or iNOS. Free Radic. Biol. Med. 35, 1217–1228 42. Suzuki, N., Kojima, H., Urano, Y., Kikuchi, K., Hirata, Y., and Nagano, T. (2002) Orthogonality of calcium concentration and ability of 4,5-diaminofluorescein to detect NO. Page 13 of 26 (page number not for citation purposes)

J. Biol. Chem. 277, 47–49 43. Zhang, X., Kim, W. S., Hatcher, N., Potgieter, K., Moroz, L. L., Gillette, R., and Sweedler, J. V. (2002) Interfering with nitric oxide measurements. 4,5-diaminofluorescein reacts with dehydroascorbic acid and ascorbic acid. J. Biol. Chem. 277, 48472–48478 44. Weinberg, J. B., Misukonis, M. A., Shami, P. J., Mason, S. N., Sauls, D. L., Dittman, W. A., Wood, E. R., Smith, G. K., McDonald, B., Bachus, K. E., et al. (1995) Human mononuclear phagocyte inducible nitric oxide synthase (iNOS): analysis of iNOS mRNA, iNOS protein, biopterin, and nitric oxide production by blood monocytes and peritoneal macrophages. Blood 86, 1184–1195 45. Vodovotz, Y., Russell, D., Xie, Q. W., Bogdan, C., and Nathan, C. (1995) Vesicle membrane association of nitric oxide synthase in primary mouse macrophages. J. Immunol. 154, 2914–2925 46. Webb, J. L., Harvey, M. W., Holden, D. W., and Evans, T. J. (2001) Macrophage nitric oxide synthase associates with cortical actin but is not recruited to phagosomes. Infect. Immun. 69, 6391–6400 47. Fang, F. C. (1997) Perspectives series: host/pathogen interactions. Mechanisms of nitric oxide-related antimicrobial activity. J. Clin. Invest. 99, 2818–2825 48. Nathan, C., and Shiloh, M. U. (2000) Reactive oxygen and nitrogen intermediates in the relationship between mammalian hosts and microbial pathogens. Proc. Natl. Acad. Sci. USA 97, 8841–8848 49. Hortelano, S., Traves, P. G., Zeini, M., Alvarez, A. M., and Bosca, L. (2003) Sustained nitric oxide delivery delays nitric oxide-dependent apoptosis in macrophages: contribution to the physiological function of activated macrophages. J. Immunol. 171, 6059–6064 50. Costantini, P., Chernyak, B. V., Petronilli, V., and Bernardi, P. (1996) Modulation of the mitochondrial permeability transition pore by pyridine nucleotides and dithiol oxidation at two separate sites. J. Biol. Chem. 271, 6746–6751 51. Hortelano, S., Alvarez, A. M., and Bosca, L. (1999) Nitric oxide induces tyrosine nitration and release of cytochrome c preceding an increase of mitochondrial transmembrane potential in macrophages. FASEB J. 13, 2311–2317 52. Coakley, R. J., Taggart, C., McElvaney, N. G., and O'Neill, S. J. (2002) Cytosolic pH and the inflammatory microenvironment modulate cell death in human neutrophils after phagocytosis. Blood 100, 3383–3391 53. Hickey, M. J., Sharkey, K. A., Sihota, E. G., Reinhardt, P. H., Macmicking, J. D., Nathan, C., and Kubes, P. (1997) Inducible nitric oxide synthase-deficient mice have enhanced leukocyte-endothelium interactions in endotoxemia. FASEB J. 11, 955–964 54. Sato, E., Simpson, K. L., Grisham, M. B., Koyama, S., and Robbins, R. A. (2000) Reactive nitrogen and oxygen species attenuate interleukin-8-induced neutrophil chemotactic activity Page 14 of 26 (page number not for citation purposes)

in vitro. J. Biol. Chem. 275, 10826–10830 55. Banick, P. D., Chen, Q., Xu, Y. A., and Thom, S. R. (1997) Nitric oxide inhibits neutrophil beta 2 integrin function by inhibiting membrane-associated cyclic GMP synthesis. J. Cell. Physiol. 172, 12–24 56. Kolb, H., and Kolb-Bachofen, V. (1998) Nitric oxide in autoimmune disease: cytotoxic or regulatory mediator? Immunol. Today 19, 556–561 57. Schaible, U. E., Winau, F., Sieling, P. A., Fischer, K., Collins, H. L., Hagens, K., Modlin, R. L., Brinkmann, V., and Kaufmann, S. H. (2003) Apoptosis facilitates antigen presentation to T lymphocytes through MHC-I and CD1 in tuberculosis. Nat. Med. 9, 1039–1046 58. Molloy, A., Laochumroonvorapong, P., and Kaplan, G. (1994) Apoptosis, but not necrosis, of infected monocytes is coupled with killing of intracellular bacillus Calmette-Guerin. J. Exp. Med. 180, 1499–1509 59. Sly, L. M., Hingley-Wilson, S. M., Reiner, N. E., and McMaster, W. R. (2003) Survival of Mycobacterium tuberculosis in host macrophages involves resistance to apoptosis dependent upon induction of antiapoptotic Bcl-2 family member Mcl-1. J. Immunol. 170, 430–437 60. Ehrt, S., Schnappinger, D., Bekiranov, S., Drenkow, J., Shi, S., Gingeras, T. R., Gaasterland, T., Schoolnik, G., and Nathan, C. (2001) Reprogramming of the macrophage transcriptome in response to interferon-gamma and Mycobacterium tuberculosis: signaling roles of nitric oxide synthase-2 and phagocyte oxidase. J. Exp. Med. 194, 1123–1140 61. Darwin, K. H., Ehrt, S., Gutierrez-Ramos, J. C., Weich, N., and Nathan, C. F. (2003) The proteasome of Mycobacterium tuberculosis is required for resistance to nitric oxide. Science 302, 1963–1966 62. Bergeron, Y., Ouellet, N., Simard, M., Olivier, M., and Bergeron, M. G. (1999) Immunomodulation of pneumococcal pulmonary infection with N(G)-monomethyl-Larginine. Antimicrob. Agents Chemother. 43, 2283–2290 63. Rijneveld, A. W., Lauw, F. N., Schultz, M. J., Florquin, S., Te Velde, A. A., Speelman, P., Van Deventer, S. J., and Van Der Poll, T. (2002) The role of interferon-gamma in murine pneumococcal pneumonia. J. Infect. Dis. 185, 91–97 Received December 29, 2003; accepted March 22, 2004.


Fig. 1


Figure 1. NO production and regulation by macrophages exposed to pneumococci. A) Human monocyte-derived macrophages (MDM) were mock-infected (Spn-) or infected with type 1 pneumococci (Spn+), and 20 h after infection cells were stained for NO using DAF-FM (left panels) or with anti-nitrotyrosine antibody (right panels). Confocal images are typical of 3 donors tested. B) NO was detected in MDM by flow cytometry after staining with DAF-FM at indicated times postinfection. Representative histograms are shown. Thick line is unstained infected MDM, and gray-filled histogram is DAF-FM stained infected MDM. Difference in mean fluorescence intensity (∆MFI) is shown in upper right corner. C) Summary of ∆MFI for 3 donors tested in duplicate, stained under same conditions as in B, after infection or mock infection. Results are means and SE. *P < 0.05 Spn- vs. Spn+, Wilcoxon signed rank test. D) Levels of iNOS protein were detected in MDM lysates 4, 12, and 20 h postinfection; 40 µg total protein loaded per lane, with actin as loading control. Results are representative of 4 donors. E) Nitrite in supernatant from MDM was detected by chemiluminescence 20 h postinfection. Results are means SE, obtained from 6 donors. *P < 0.05 Spn- vs. Spn+, **P < 0.01 Spn+ vs. Spn+ 1400W+, Friedman test with Dunn’s multiple comparison test.


Fig. 2


Figure 2. NO production in macrophages does not localize to lysosomes, bacteria, or mitochondria. MDM were mock-infected (Spn-) or infected with type 1 pneumococci (Spn+) and NO detected by DAF-FM. NO production resulted in a green staining pattern that became more punctate with time postinfection (left panels). Cells were costained with markers of specific organelles or bacteria, as depicted in red (central panels). Overlay images showing both red and green staining demonstrate a lack of colocalization as represented by an absence of yellow color (right panels). Arrowheads show representative areas of DAF-FM staining and arrows show areas of staining of the labeled organelle in the overlay image. Bacteria are indicated with a double arrow. MDM were costained 14 h after infection with LysoTracker to detect lysosomes (A), propidium iodide to stain bacteria and nuclei 16 h after infection (B), or MitoTracker to stain mitochondria 16 h after infection (C). MDM were imaged by fluorescence microscopy, and images are representative of 3 experiments.


Fig. 3

Figure 3. NO contributes to killing of intracellular pneumococci. A) Number of bacteria internalized by MDM 4 h postinfection with type 1 pneumococci in the absence (-) or presence (+) of the iNOS inhibitor 1400W. Results are means and SE obtained from 4 donors. B) Intracellular killing assays were performed in MDM 6 and 16 h postinfection with pneumococci. Results are means and SE obtained from the same 4 donors during the same experiments as in A. *P < 0.05, Spn+ vs. Spn+ 1400W+, Wilcoxon signed rank test.


Fig. 4

Figure 4. NO production is associated with apoptosis induction in macrophages exposed to pneumococci. A) MDM were mock-infected (Spn-) or infected with type 1 pneumococci (Spn+) for 20 h in the presence (+) or absence (-) of 1400W, and apoptosis was detected by TUNEL with propidium iodide counterstaining. Images were recorded by fluorescence microscopy and green/yellow cells are TUNEL+. B) Percentage of TUNEL+ cells was quantified by fluorescence microscopy. Results are means and SE of percentage apoptosis for duplicate samples obtained from 5 donors. **P < 0.01 Spn+ vs. Spn+ 1400W+ Mann-Whitney U test. C) Nuclear features of apoptosis recorded in MDM 20 h postinfection in the presence (+) or absence (-) of 1400W or L-NMA. Results are means and SE of percent apoptosis for duplicate samples obtained from four donors. *P < 0.05 Spn- vs. Spn+, Spn+ vs. Spn+ 1400W+ and Spn+ vs. Spn+ L-NMA+, Wilcoxon signed rank test. D) Nuclear features of apoptosis recorded in MDM 20 h postinfection in the presence or absence of indicated concentrations of NOC-18. Results are means and SE of percent apoptosis obtained from duplicate samples from 3 donors. *P < 0.05 Spn+ vs. Spn+ 100 µg/ml NOC-18+, Kruskal-Wallis test with Dunn’s multiple comparison.


Fig. 5


Figure 5. NO production during pneumococcal infection induces mitochondrial membrane permeabilization. A) MDM were mock-infected (Spn-) or infected with type 1 pneumococci (Spn+) for 16 h in presence (+) or absence (-) of 1400W and stained with JC-1. Loss of ∆ψm is represented by a decrease in punctate red staining. Results represent at least 5 separate experiments. B) Loss of ∆ψm in MDM was assessed 16 h postinfection by flow cytometry after staining with JC-1. Representative histograms from 1 donor are shown. Percentages represent number of cells with loss of ∆ψm. C) Percentage of cells with loss of ∆ψm from 6 donors under same conditions as B; *P < 0.05 Spn+ vs. Spn+ 1400W+, Wilcoxon signed rank test. D) Western blots of the cytoplasmic fraction obtained from MDM 20 h after infection in the presence (+) or absence (-) of 1400 W probed with anti-cytochrome c antibody; 20 µg protein loaded per lane, with reprobing with actin to confirm equal protein loading. Data represent 3 independent experiments. E) Western blot of total protein obtained from MDM 20 h after infection probed with anti-Mcl-1; 25 µg protein loaded per lane, with reprobing with actin to confirm equal protein loading. Results represent 3 independent experiments.


Fig. 6

Figure 6. Mitochondrial membrane permeabilization is a critical feature of pneumococcal-associated macrophage apoptosis. Nuclear features of apoptosis were recorded in MDM 20 h after infection with type 1 pneumococci (Spn+) or mock-infection (Spn-) in presence (+) or absence (-) of the indicated doses of bongkrekic acid (BA). Results are means and SE of percent apoptosis obtained from 3 donors tested in duplicate. ***P < 0.001 Spn- vs. Spn+, *P < 0.05 Spn+ vs. Spn+ with 50 µM BA, Friedman test with Dunn’s multiple comparison test.


Fig. 7

Figure 7. iNOS inhibition switches macrophage death during pneumococcal infection from apoptosis to necrosis. A) MDM were mock-infected or infected with type 1 pneumococci (Spn) for 20 h in the presence (+) or absence (-) of 1400 W, and cell density was assessed. Results are means and SE for 4 donors tested in duplicate. *P < 0.05 Spn- vs. Spn+, Spn- 1400 W+ vs. Spn+ 1400W+, Wilcoxon signed rank test. B) Caspase activation was measured by FITC-VADFMK staining (FITC-VAD) and necrosis by propidium iodide staining (PI) in MDM 16 h postinfection. Percent events in each quadrant are shown on the right. Results represent 3 independent experiments.


Fig. 8

Figure 8. Pneumolysin triggers macrophage apoptosis and NO production. A) MDM were incubated for 20 h in varying concentrations of pneumolysin, and nuclear features of apoptosis were recorded. Results are means and SE of percent apoptosis for 5 donors tested in duplicate. *P < 0.05, 0 vs. 10 µg/ml pneumolysin and 0.1 vs. 10 µg/ml pneumolysin, Friedman test with Dunn’s multiple comparison test. B) Nuclear features of apoptosis in MDM 20 h after mock-infection (MI), infection with pneumolysin deficient type 2 pneumococci (PLY-), infection with parental type 2 pneumococci (D39), or infection with pneumolysin deficient type 2 pneumococci in the presence of 1 µg/ml of exogenous pneumolysin (PLY- +PLY). Results are means and SE of percent apoptosis for 6 donors tested in duplicate. *P < 0.05 D39 vs. PLY- PLY- vs. PLY-+PLY, Wilcoxon signed ranks test. C) Western blot of the cytosolic fraction obtained from MDM 20 h after infection probed with anti-cytochrome c; 20 µg protein loaded per lane, with reprobing with actin to confirm equal protein loading. Results represent 3 individual experiments. D) MDM were stained with DAF-FM to detect NO 20 h after infection and analysed by flow cytometry. Results are means and SE of change in median fluorescence intensity versus unstained samples (∆MFI) for 5 donors. *P < 0.05 D39 vs. PLY-, Wilcoxon signed ranks test.
