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The Journal of Immunology, 2000, 165: 3923-3933.
Copyright © 2000 by The American Association of Immunologists

Group B Streptococcus Induces Apoptosis in Macrophages1

Katia Fettucciari*, Emanuela Rosati*, Lucia Scaringi*, Paola Cornacchione*, Graziella Migliorati{dagger}, Rita Sabatini*, Ilaria Fetriconi*, Ruggero Rossi* and Pierfrancesco Marconi2,*

* General Pathology and Immunology Section, {dagger} Toxicology Pharmacology and Chemotherapy Section, Department of Clinical and Experimental Medicine, University of Perugia, Perugia, Italy


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Group B Streptococcus (GBS) is a pathogen that has developed some strategies to resist host immune defenses. Because phagocytic killing is an important pathogenetic mechanism for bacteria, we investigated whether GBS induces apoptosis in murine macrophages. GBS type III strain COH31 r/s (GBS-III) first causes a defect in cell membrane permeability, then at 24 h, apoptosis. Apoptosis was confirmed by several techniques based on morphological changes and DNA fragmentation. Cytochalasin D does not affect apoptosis, suggesting that GBS-III needs not be within the macrophage cytoplasm to promote apoptosis. Inhibition of host protein synthesis prevents apoptosis, whereas inhibition of caspase-1 or -3, does not. Therefore, GBS can trigger an apoptotic pathway independent of caspase-1 and -3, but dependent on protein synthesis. Inhibition of apoptosis by EGTA and PMA, and enhancement of apoptosis by calphostin C and GF109203X suggests that an increase in the cytosolic calcium level and protein kinase C activity status are important in GBS-induced apoptosis. Neither alteration of plasma membrane permeability nor apoptosis were induced by GBS grown in conditions impeding hemolysin expression or when we used dipalmitoylphosphatidylcholine, which inhibited GBS ß-hemolytic activity, suggesting that GBS ß-hemolysin could be involved in apoptosis. ß-Hemolysin, by causing membrane permeability defects, could allow calcium influx, which initiates macrophage apoptosis. GBS also induces apoptosis in human monocytes but not in tumor lines demonstrating the specificity of its activity. This study suggests that induction of macrophage apoptosis by GBS is a novel strategy to overcome host immune defenses.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Group B Streptococcus (GBS)3 is a pathogen that causes serious neonatal infections (1). It resists phagocytosis by macrophages and polymorphonuclear cells by means of an antiphagocytic capsule (1, 2, 3) and, like an intracellular microorganism, can survive for some time inside respiratory epithelial cells, endothelial cells (4, 5, 6), and macrophages (7, 8). The recent discovery that GBS can kill endothelial cells, epithelial cells, and fibroblasts by ß-hemolysin (9, 10, 11) suggests that this microorganism may also have evolved an anti-host strategy based on the killing of host immune cells. It is well known that several microbial pathogens kill immune cells by apoptosis or necrosis (12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23). In particular, it has been demonstrated that apoptosis plays an important role in various infectious diseases (24, 25, 26, 27). Moreover, several microorganisms have been shown to cause apoptosis in macrophages to counteract host immune defenses (13, 14, 15, 16, 17, 18, 19, 20). The ability of pathogens to promote apoptosis may be important for the initiation of infection, bacterial survival, and escape from the host immune response. In fact, because apoptosis occurs without the release of cellular components, it does not usually lead to inflammation (28, 29, 30). Therefore, apoptosis may be advantageous for the pathogen because it might avoid the triggering and recruitment of non specific host defense mechanisms. Furthermore, macrophage death could also contribute to delaying or hindering the development of a specific immune response.

Apoptosis is characterized by a number of biochemical events, including protein kinase C (PKC) activity, cytoplasmic calcium (Ca2+) increase, and caspase activation (29, 31, 32, 33, 34, 35). Because an understanding of the apoptosis mechanism induced by bacteria could be important for the management of infectious diseases, we investigated the ability of GBS to induce apoptosis in macrophages and the mechanisms involved.

This study shows that GBS induces apoptosis in murine macrophages and that inhibition of Ca2+ influx and PKC activation counter GBS-induced macrophage apoptosis, whereas caspase inhibition does not. Inhibition of apoptosis both by growing GBS in conditions, which abolish the synthesis of GBS ß-hemolysin, and by dipalmitoylphosphatidylcholine (DPPC), an inhibitor of ß-hemolytic activity, suggests that ß-hemolysin could be involved in the induction of apoptosis. However, apoptosis induction by GBS does not seem a general mechanism because GBS causes apoptosis also in human monocytes but not in other cell types.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals

Outbred CD-1 mice of both sexes, 8–10 wk old, were obtained from Charles River Breeding Laboratories (Calco, Milan, Italy).

Chemicals

Cycloheximide (CHX), PMA, cytochalasin D (Cyt D), actinomycin D (Act D), EGTA, calphostin C, GF109203X, DPPC, and propidium iodide (PI) were from Sigma (St. Louis, MO). The inhibitor of caspase-1 like protease inhibitor V (ZVAD.fmk) and the inhibitor of CPP32/apopain, caspase-3 (DEVD-CHO) were from Calbiochem (La Jolla, CA).

Microorganisms

GBS type III, strain COH31 r/s (GBS-III), clinically isolated from a diabetic foot ulcer of an adult, rendered resistant to rifampicin and streptomycin, GBS type III, strain COH 1 and GBS type VI, strain 118754 were kindly provided by Dr. M. Wessel (Channing Laboratory, Boston, MA). GBS type Ia mouse passed prototype stain 090 was kindly provided by Dr. J. Jelinkova (Institute of Hygiene and Epidemiology, Prague, Czech Republic). All strains were grown in Todd-Hewitt broth (THB; Unipath, Milan, Italy) at 37°C, and aliquots were stored at -70°C until used.

For assays, the microorganisms were grown in THB overnight and then washed and adjusted photometrically (600 nm) to the desired number of CFU per ml. The concentration and purity of inoculum was confirmed by quantitative culture on Islam agar (Unipath) plates containing 5% heat-inactivated horse serum. For some experiments GBS-III were grown for 18 h in THB in the presence of 10 mg/ml glucose (gGBS-III), conditions not allowing hemolytic activity expression (36), and then treated as described above.

Peritoneal murine macrophages, human blood monocytes, and tumor cell lines

Mice were injected i.p. with 1 ml of a 10% solution of Bacto Brewer Thioglycollate medium (thioglycollate broth, Difco, Detroit, MI). After 4 days, peritoneal exudate cells were harvested by washing the peritoneal cavity with 10 ml cold RPMI 1640 medium containing 5 U/ml heparin and aspirating the exudate with a syringe. The cells were washed three times in cold antibiotic-free RPMI 1640 with 5% FCS (complete medium) and cell viability was evaluated by trypan blue exclusion method.

Peripheral blood monocytes were isolated from human healthy donors by standard Ficoll-Hypaque (Sigma) gradient centrifugation and monocytes separated from lymphocytes by centrifugation (650 x g) for 20 min over a 35/51% Percoll gradient (Pharmacia, Piscataway, NJ). Monocytes recovered from the interphase were washed three times in RPMI, and cell viability was evaluated by trypan blue exclusion method. The cells were >95% pure monocytes as determined by immunofluorescence staining with the CD14 (PharMingen, San Diego, CA) Ab and flow cytometry analysis. The human T cell lymphoma HUT 78, human T cell leukemia JURKAT, human B cell lymphoma RAJI, murine T cell lymphoma YAC-1, and murine mast cells P-815 were maintained in RPMI with 10% FCS.

Infection procedure

The macrophages, human monocytes, and tumor cell lines, adjusted to a concentration of 1 x 106cells/ml in complete medium, were infected in 12- x 75-mm polypropylene tubes with GBS-III at a cell:microorganism ratio of 1:100. For preliminary experiments, we used macrophages infected for 0.5, 1, 1.5, and 2 h and macrophages infected for 2 h washed and reincubated for 12, 24, and 48 h in complete medium containing 100 U/ml penicillin and 100 µg/ml gentamicin. In the following experiments, macrophages, human monocytes, or tumor cell lines infected for 2 h and reincubated for 12 and 24 h in medium containing antibiotics were used. Control cells were incubated in medium for the same times.

The infection of macrophages with GBS type III, strain COH 1, GBS type Ia, strain 090, GBS type VI, strain 118754 and gGBS-III was performed as described above.

For experiments with Cyt D and caspase and PKC inhibitors, Cyt D (1 µg/ml), ZVAD.fmk (50 µM), DEVD-CHO (50 µM), GF109203X (1 µM), or calphostin C (0.5 µM) was added to macrophages 30 min before infection with GBS-III and maintained during the course of the experiments. In the experiments with CHX (50 µg/ml), the inhibitor was added to macrophages 30 min before infection with GBS-III and maintained during 2 h of infection, then removed because a prolonged exposure of macrophages to CHX is cytotoxic. Macrophages treated with inhibitors but not infected were used as controls.

For experiments with EGTA and PMA, 1 mM or 1 µg/ml, respectively, were added to macrophages during the 2-h infection, and 0.5 mM EGTA was maintained throughout the course of the experiment, whereas PMA was removed because a prolonged treatment with PMA specifically depletes PKC activity.

For DPPC experiments, sonicated DPPC (1 min, 30 W) was added at concentrations of 1, 0.5, and 0.25 mg/ml to macrophages during the 2-h infection and then removed. Macrophages treated with DPPC but not infected were used as controls.

PI uptake assay

At different time points, infected cells and controls were washed, adjusted to 1 x 106/ml in PBS containing PI (5 µg/ml, Sigma), incubated at 23°C for 5 min, and analyzed on a FACScan flow cytometer (Becton Dickinson). PI penetrates and intercalates into the DNA of cells which have lost membrane integrity, causing them to fluoresce red when activated with UV light (21).

Assay for hemolytic activity

To quantify GBS hemolytic activity of whole bacteria, a modified Marchlewicz and Ducan method was used (36). GBS-III was grown in THB, washed, and adjusted to 109 CFU/ml in PBS. In a 96-well conical bottom microtiter plate, 100 µl (108 CFU) of the bacterial suspension was added to the first well, and serial 2-fold dilutions in PBS were made across the plate, each in a final volume of 100 µl. An equal volume of 1% SRBC in PBS was added to each well, and the plate incubated at 37°C in 5% CO2 for 2 h. SRBC incubated in PBS alone and SRBC incubated with 0.1% SDS were used as negative and positive controls, respectively. After 2 h, the plates were spun at 3000 x g for 10 min, 100 µl of supernatant were transferred to a fresh plate, and OD542 nm was measured. Hemolytic titer was determined as the inverse of the greatest dilution producing 50% hemoglobin release compared with SDS control. The GBS inoculum size and assay times were the same as in the infection procedure.

The hemolytic assay with gGBS and with different GBS strains (109 CFU/ml) was performed as described above for GBS-III.

For DPPC experiments, sonicated DPPC (1 min, 30 W) was added at concentrations of 1, 0.5, and 0.25 mg/ml to wells containing SRBC and GBS-III. The assay was performed as described above. SRBC incubated with the different doses of DPPC without GBS were used as negative controls.

DNA labeling and flow cytometry analysis

At different time points, infected cells and controls were recovered for detection of apoptosis. Macrophages incubated with 1 µg/ml Act D were used as a positive control for apoptosis (37). Macrophages heated at 42°C for 1 h and then incubated for different times at 37°C or macrophages incubated with 0.2% saponin were used as control for necrosis (38). After washing, the 200 x g centrifuged cell pellets were resuspended in 1 ml of hypotonic fluorochrome solution (PI 50 µg/ml in 0.1% sodium citrate plus 0.1% Triton X-100; Sigma). The samples were placed overnight in the dark at 4°C, and the PI fluorescence of individual nuclei measured using a FACScan flow cytometer as described by Nicoletti et al. (39). The data were processed by a Hewlett Packard computer and analyzed with Lysis software (Becton Dickinson).

Electron microscopy

At 24 h after addition of antibiotics, infected macrophages and controls were washed with cacodylate buffer (pH 7.4) and fixed for 2 h in 2.5% glutaraldehyde in 0.1 M cacodilate buffer (pH 7.4). After washing in buffer, the cells were postfixed in 2% osmium tetroxide plus 1% potassium ferricyanide in cacodylate buffer, dehydrated through a graded alcohol series, embedded in Epon, thin sectioned, stained with uranyl acetate and lead citrate, and observed by electron microscopy. Macrophages incubated with 1 µg/ml Act D for 24 h were used as a positive control for apoptosis (37).

DNA fragmentation assay

DNA was isolated from controls and infected macrophages 24 h after addition of antibiotics, from Act D-treated macrophages used as positive control for apoptosis (37), and from heat-treated macrophages, used as positive control for necrosis (38). DNA fragmentation was assessed according to the method described elsewhere (39). Briefly, cells (4 x 106 macrophages) were collected by centrifugation at 200 x g for 10 min and dissolved in hypotonic lysing buffer (100 mM NaCl, 10 mM Tris, 1 mM EDTA, 1% SDS, 200 mg/ml proteinase K, pH 7.5). The lysates were deproteinized by extraction, twice with phenol-chloroform-isoamyl alcohol (25:24:1) and once with chloroform-isoamyl alcohol (24:1) then precipitated overnight at -20°C in 2 vol ethanol in the presence of 0.3 M sodium acetate and recovered by centrifugation. Pellets were air dried, and resuspended in 10 mM Tris-HCl (pH 8.0) 1 mM EDTA (TE buffer) at 4°C. The DNA solution was then analyzed in a 2% agarose gel stained with ethidium bromide.

Nick-end labeling DNA

The frequency of apoptotic cells in GBS-infected cells was also quantified by the TUNEL technique using an in situ cell death detection kit (Boehringer Mannheim, Mannheim, Germany) (40). The TUNEL reaction preferentially labels DNA strand breaks generated during apoptosis. Labeling was performed according to the manufacturer’s instructions. Briefly, 1 x 106 cells were fixed in 2% paraformaldehyde in 1x PBS and then permeabilized in 0.1% triton, 0.1% citrate for 2 min on ice. The 3'-OH termini of internucleosomal DNA strand breaks were labeled with TUNEL reaction mixture for 1 h at 37°C. Cells were then analyzed by flow cytometry on a Becton Dickinson FACScan.

Statistical analysis

Experiments of PI uptake, trypan blue exclusion methods, and flow cytometric DNA analysis in the presence of inhibitors were repeated six times. Data are presented as the means ± SD of six independent experiments performed in triplicate. The data for each experiment were analyzed by Student’s t test. The electron microscopy analysis and analysis of DNA fragmentation by flow cytometry and agarose gel electrophoresis were repeated six times and data reported are of a typical experiment.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
GBS-III induces macrophage plasma membrane permeability

In initial experiments, the effect of GBS-III on the plasma membrane permeability of thioglycollate-elicited peritoneal murine macrophages was analyzed at different times after infection, by PI uptake assay. GBS-III was able to induce alterations in plasma membrane permeability (Fig. 1GoA). The percentage of macrophages with permeabilized membranes, significantly increased to about 72% PI+ cells at 2 h after infection, remained around 70% during the following 24 h of culture and then decreased to about 32% at 48 h after infection (Fig. 1GoA). To determine whether the number of macrophages with increased membrane permeability decreases in time as a result of cell death, the total cell number at different times after infection was evaluated by trypan blue exclusion assay. Results indicate that GBS-III induces macrophage death because it causes a 70% decrease in the total cell number, evaluated at 48 h after infection (Fig. 1GoB).



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  		FIGURE 1. Effect of GBS on macrophage membrane permeability and cell death. Macrophages infected with GBS-III and recovered at different times after infection. At the indicated time points, the percentage of PI+ cells by PI uptake assay (A), or the total cell number by trypan blue assay (B), was determined. The data are means ± SD of six experiments performed in triplicate. *, p < 0.01 (GBS-infected macrophages vs control macrophages) according to Student’s t test.

 
Microbial factors responsible for GBS-III alterations in macrophage plasma membrane permeability

To identify the microbial factors responsible for plasma membrane defects, we first examined the role of soluble factors released by GBS-III in the culture medium. To this end, macrophages were incubated with GBS-III in contiguous medium separated by a 0.45-µm pore size membrane of cell culture insert and PI uptake was performed at different times after infection. Under these conditions, no alterations in membrane permeability were observed at all times examined (Fig. 2GoA). These results suggest that macrophage plasma membrane permeability defects depend on GBS factor(s), which is/are insoluble or unstable in supernatants, and require GBS-III-macrophage contact.



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  		FIGURE 2. Effect of GBS ß-hemolysin on macrophage membrane permeability. The percentage of PI+ cells was determined at different times by PI+ uptake assay. The data are means ± SD of six experiments performed in triplicate. A, Macrophages infected with GBS-III, gGBS-III, or GBS-III on a 0.45-µm pore filter placed above macrophages, and recovered at different times after infection. B, Macrophages infected for 2 h with GBS-III, in the absence or presence of DPPC (1, 0.5, and 0.25 mg/ml) and recovered at different times after infection. Treatment of control macrophages with each concentration of DPPC for 2 h did not determine loss of cell viability at all times examined. *, p < 0.01 (GBS-III-infected macrophages treated with DPPC vs untreated GBS-III-infected macrophages) according to Student’ s t test.

 
It is known that GBS has a ß-hemolysin bound to the cell surface, which is highly unstable when released in culture supernatants (36, 41). However, it has been recently demonstrated that the ß-hemolysin is active against the membrane of some eukaryotic cells (9, 10, 11). To investigate whether GBS-III ß-hemolysin can alter macrophage membrane permeability, we first used gGBS-III, grown for 18 h in the presence of 10 mg/ml glucose, a condition that abolishes GBS ß-hemolysin synthesis (36). Results of PI uptake assay indicate that gGBS-III did not cause changes in the plasma membrane permeability. Indeed the percentage of PI+ macrophages was similar in both controls and gGBS-III infected macrophages at all times examined (Fig. 2GoA). Loss of ß-hemolysin synthesis, due to growing GBS in presence of glucose, was confirmed by evaluating hemolytic activity of gGBS-III against SRBC (Table IGo).


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  		Table I. Loss of ß-hemolytic activity by growing GBS-III in the presence of glucose

 
Because it has been reported that some phospholipids such as DPPC inhibited GBS ß-hemolytic activity (10, 36), we analyzed the effect of DPPC on GBS-induced plasma membrane permeability defects by PI uptake assay. It was found that DPPC inhibited the GBS-III-induced membrane permeability defects in a dose-dependent manner (Fig. 2GoB). With 1 mg/ml of DPPC, plasma membrane permeability defects were almost completely inhibited at all times examined (Fig. 2GoB). Instead, with 0.5 and 0.25 mg/ml DPPC, the plasma membrane permeability was inhibited by 74 and 20%, respectively, at all times tested (Fig. 2GoB).

DPPC inhibition of GBS-III ß-hemolytic activity was confirmed by SRBC hemolysis (Table IIGo).


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  		Table II. Inhibition of GBS-III hemolytic activity by DPPC

 
The results suggest that the effect of GBS on macrophage plasma membrane permeability is mediated by ß-hemolysin.

GBS-III promotes macrophage apoptosis

To characterize the nature of cell death caused by GBS-III, we infected macrophages and, at different times after infection, measured changes in DNA content by flow cytometry (39). The flow cytometry profile of PI-stained macrophages at 2 h after GBS-III infection showed a single diploid DNA peak and the percentage of diploid nuclei was similar to that of uninfected macrophages, whereas at 12 h a hypodiploid DNA peak, corresponding to 26.5% apoptotic cells appeared only in GBS-III infected macrophages. At 24 h after infection, apoptotic cells were about 82.5% (Fig. 3Go). A similar percentage of apoptotic cells (79%) was observed in macrophages treated for 24 h with Act D, a potent inducer of apoptosis (Fig. 3Go) (37). On the contrary, no hypodiploid DNA peaks were observed in macrophages treated with necrotic stimuli (38) such as heating at 42°C for 1 h (Fig. 3Go) or treatment with 0.2% saponin (data not shown).



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  		FIGURE 3. DNA fluorescence flow cytometric profiles of PI-stained macrophages. Control, GBS-III infected macrophages, macrophages incubated with 1 µg/ml Act D (positive control for apoptosis), and macrophages heated at 42°C for 1 h (positive control for necrosis) recovered at different times after infection. Apoptosis was determined at the time points indicated measuring the percentage of hypodiploid nuclei at flow cytometry. Percentage numbers of hypodiploid nuclei are reported for each time and condition. One experiment, representative of six, is shown.

 
Morphology and DNA fragmentation of macrophages infected with GBS-III

To confirm that cell death measured by flow cytometry corresponded to apoptosis, we analyzed by electron microscopy the morphology of macrophages 24 h after GBS-III infection. The majority of macrophages (about 80%) at 24 h after infection showed clear signs of apoptosis: cell shrinkage, chromatin condensation, nuclear fragmentation, cytoplasmic vacuolization, and preservation of organelle structure (Fig. 4GoA). A similar morphology was observed in macrophages treated for 24 h with Act D (Fig. 4GoB). On the contrary, uninfected control macrophages had a normal appearance (Fig. 4GoC).



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  		FIGURE 4. Electron micrographs of macrophages infected with GBS. A, Macrophages at 24 h after infection with GBS-III showing apoptotic morphology: chromatin condensation and nuclear fragmentation, cytoplasmic vacuolization and maintenance of organelle structure; magnification x2,390; B, macrophages incubated with Act D (1 µg/ml for 24 h) with apoptotic morphology similar to that of GBS-III infected macrophages; magnification x2,390; C, control macrophages incubated in medium for 24 h showing normal morphology; magnification x2,390.

 
GBS-III-induced macrophage apoptosis was also evaluated by agarose gel electrophoresis of DNA. A characteristic internucleosomal banding pattern was detected in DNA extracted from macrophages at 24 h after GBS-III infection but not in DNA from control cells (Fig. 5GoA). A pattern of DNA fragmentation similar to that of infected macrophages was observed in Act D-treated cells (Fig. 5GoA). Necrotic stimuli such as heating at 42°C for 1 h (Fig. 5GoA) or 0.2% saponin (data not shown) did not lead to DNA fragmentation.



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  		FIGURE 5. DNA fragmentation in GBS-infected macrophages. A, Agarose gel electrophoresis. Total cellular DNA was isolated at 24 h from control, GBS-III-infected, Act D-treated (1 µg/ml for 24 h), and heat-treated (42°C for 1 h) macrophages. Right lane shows m.w. marker, pBR328/BglI digest + pBR328/HinfI digest. The DNA samples were analyzed by electrophoresis in 2% agarose gel with ethidium bromide. Only Act D-treated cells and GBS-III-infected macrophages generated ladders of 200 bp, characteristic of apoptosis. One experiment, representative of six, is shown. B, TUNEL reaction. Control and GBS-III-infected macrophages were processed at 24 h for detection of DNA strand breaks by TUNEL and analyzed at flow cytometry. The histograms show the fluorescein dUTP incorporation. Percentage numbers of cells that had incorporated dUTP are reported for each condition. One experiment, representative of six, is shown.

 
Detection of apoptotic cells using the TUNEL technique (40) closely correlates with results obtained using PI staining of nuclei. In fact, at 24 h after infection, about 76% of GBS-III-infected macrophages had incorporated large quantities of fluorescein dUTP after incubation with TdT (Fig. 5GoB). Because DNA fragments detected by TUNEL assay were specific for apoptosis, these data confirm the results obtained with PI staining of host nuclei indicating that GBS-infected macrophages underwent cell death by apoptosis.

Role of ß-hemolysin on GBS-III-induced macrophage apoptosis

Because the induction of apoptosis by several microorganisms depends on their ability to enter cell cytosol (13, 15, 17) we analyzed whether bacterial internalization was also a requirement for GBS-III-induced apoptosis. Macrophages were pretreated for 30 min with Cyt D, a drug that inhibits actin polymerization, thereby preventing phagocytosis. The cells were then infected for 2 h with GBS-III, and at different times after infection, recovered for PI staining and flow cytometry analysis. As shown in Fig. 6GoA, Cyt D did not protect macrophages from apoptosis. These results indicate that GBS-III can trigger macrophage apoptosis from an extracellular localization.



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  		FIGURE 6. Effect of ß-hemolysin on GBS-induced macrophage apoptosis. Macrophage apoptosis measured at different times, evaluating the percentage of hypodiploid nuclei at flow cytometry. The data are means ± SD of six experiments performed in triplicate. A, Macrophages pretreated for 30 min in the presence or absence of Cyt D (1 µg/ml) infected with GBS-III and recovered at different times after infection. B, Macrophages infected with gGBS-III and recovered at different times after infection. C, Macrophage infected by placing the GBS-III on a 0.45-µm pore filter above macrophages and recovered at different times after infection. D, Macrophages infected with GBS-III in the absence or presence of DPPC (1, 0.5, and 0.25 mg/ml) and recovered at different times after infection. Treatment of control macrophages with each concentration of DPPC for 2 h did not induce apoptosis at all times examined. *, p < 0.01 (GBS-III infected macrophages treated with DPPC vs untreated GBS-III infected macrophages) according to Student’ s t test.

 
In a further series of experiments, we investigated the possible microbial factors involved in GBS-III-induced apoptosis. GBS has a ß-hemolysin (36), which like pore-forming proteins, causes membrane permeability defects. Because it is generally assumed that pore-forming proteins can generate alterations in membrane permeability and also induce apoptosis (12, 29), we examined the possibility that GBS-III could also cause macrophage apoptosis indirectly through ß-hemolysin. First, the changes in DNA content in macrophages incubated with gGBS-III, which had lost ß-hemolytic activity, were measured by flow cytometry. The gGBS-III did not induce macrophage apoptosis at all times examined (Fig. 6GoB). Because ß-hemolysin is firmly bound to the cell surface (41) and is unstable when released in the supernatants (36), we tested the effect of GBS-III in conditions that did not allow GBS-macrophage interaction. Macrophages were incubated with GBS-III in contiguous media, separated by a 0.45-µm pore size membrane filter. Nor in this case was there GBS-III-induced apoptosis because the percentage of apoptotic cells of infected and uninfected macrophages was similar (Fig. 6GoC).

To gain further insight into the possible role of ß-hemolysin in macrophage apoptosis, we examined whether GBS-induced apoptosis was affected by DPPC, which is known to inhibit GBS ß-hemolytic activity (Refs. 10, 36 , and Table IIGo). DPPC inhibited GBS-III induced apoptosis in a dose-dependent manner (Fig. 6GoD). DPPC at 1 mg/ml and 0.5 mg/ml (added during 2 h of infection) inhibited apoptosis, by about 85 and 70%, respectively, both at 12 h and 24 h after infection, evaluated by cytofluorometric analysis. Instead, 0.25 mg/ml DPPC, inhibited macrophage apoptosis by about 20% (Fig. 6GoD).

To further test the hypothesis that ß-hemolysin could be involved in macrophage apoptosis, we used four GBS strains varying in ß-hemolysin expression (10). The results show that these strains induced apoptosis in close correlation to their ability to lyse SRBC (Table IIIGo). In fact, with weakly hemolytic GBS type III strain COH 1 (hemolytic titer 16) there was 40% macrophage apoptosis, and with GBS type VI strain 118754 (hemolytic titer 32), the percentage of apoptosis was 60%, with GBS type Ia strain 090 (hemolytic titer 64), and GBS-III (hemolytic titer 64) the percentage of apoptosis reached a value >80% (Table IIIGo).


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  		Table III. Correlation of hemolytic titer of different GBS strains with macrophage apoptosis

 
The results suggest that GBS ß-hemolysin could be involved in macrophage apoptosis.

GBS-III induces plasma membrane permeability defects in human monocytes and tumor lines but apoptosis only in human monocytes

To determine whether GBS-induced alterations in plasma membrane permeability and apoptosis were specific for murine macrophages, we studied the effects of GBS on human monocytes and on different human and murine tumor lines. Human monocytes, HUT 78, RAJI, JURKAT, YAC-1, and P-815 cells were infected with GBS-III for 2 h. At different times after infection, alterations in plasma membrane permeability and total cell number were evaluated by PI uptake assay and trypan blue exclusion method, respectively. The percentage of PI+ cells in both human monocytes (Fig. 7GoA) and tumor lines (Fig. 7Go, B–F) was about 80% at 2 h after infection, remained around this value during the following 24 h and then decreased to about 30% at 48 h (Fig. 7Go). Therefore, GBS-III causes alterations in plasma membrane permeability in all cell types examined as in thioglycollate-elicited peritoneal macrophages. However, the total number of GBS-III infected tumor cell lines, evaluated by trypan blue assay, decreased by about 50% at 2 h after infection and continued to decrease to 70% at 12 h (Fig. 7Go, B–F), whereas the total number of GBS-III-infected human monocytes decreased by about 70% only at 48 h after infection (Fig. 7GoA).



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  		FIGURE 7. Effect of GBS on plasma membrane permeability and cell death of human monocytes and tumor lines. Human monocytes (A), HUT 78 (B), RAJI (C), JURKAT (D), YAC-1 (E), and P-815 (F) cells infected with GBS-III were recovered at different times after infection and the percentage of PI+ cells (lines: {square}, control; {blacktriangleup}, GBS-III) by PI uptake assay or the total cell number (hystograms: {square}, control; {blacksquare}, GBS-III) by trypan blue assay was determined. The data are means ± SD of six experiments performed in triplicate. *, p < 0.01 (GBS-III-infected cells vs control cells) according to Student’ s t test.

 
To determine whether GBS-III induced apoptosis, human monocytes, HUT 78, RAJI, JURKAT, YAC-1, and P-815 cells were infected with GBS-III and at different times recovered for PI staining of nuclei and flow cytometry analysis. GBS-III induced apoptosis in human monocytes (Fig. 8GoA). In fact, a significant percentage of apoptotic cells (about 25%) was revealed at 12 h after infection and continued to increase to about 80% at 24 h. (Fig. 8GoA). On the contrary, no apoptosis was observed in any tumor cell line, either in GBS-III infected or uninfected at all times examined (Fig. 8Go, B–F). TUNEL technique confirmed the results obtained using PI staining of DNA. There was incorporation of large quantities of fluorescein dUTP after incubation with TdT in GBS-infected human monocytes corresponding to 80% of apoptotic cells, at 24 h after infection but not in GBS-infected tumor lines at all times examined (data not shown).



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  		FIGURE 8. GBS induces apoptosis in human monocytes but not in tumor lines. Human monocytes (A), HUT 78 (B), RAJI (C), JURKAT (D), YAC-1 (E), and P-815 (F) cells infected with GBS-III ({blacksquare}) or uninfected ({diamond}) were recovered at different times after infection and apoptosis measured evaluating the percentage of hypodiploid nuclei at flow cytometry. The data are means ± SD of six experiments performed in triplicate. *, p < 0.01 (GBS-III-infected cells vs control cells) according to Student’ s t test.

 
All these findings indicate that GBS-III causes alterations in plasma membrane permeability both in human monocytes and tumor cell lines but induced apoptosis only in human monocytes.

Inhibition of GBS-III-induced apoptosis

Macromolecular synthesis is required in several types of apoptosis (29). We evaluated whether CHX, an inhibitor of cell protein synthesis, had any effect on GBS-III-induced macrophage apoptosis. Treatment with 50 µg/ml CHX 30 min before and during the 2 h of infection blocked GBS-III-induced macrophage apoptosis. Table IVGo shows that at 24 h after infection, apoptosis was inhibited in CHX-treated macrophages, demonstrating that GBS-III-induced apoptosis depended on de novo protein synthesis.


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  		Table IV. Effect of inhibitors on GBS-III-induced macrophage apoptosis

 
Apoptosis requires tightly regulated death pathways including activation of caspases (33, 34, 35). To determine the role of caspases in GBS-III-induced apoptosis, the possible effect of membrane permeable irreversible inhibitor caspase-1 (ZVAD.fmk) and reversible inhibitor caspase-3 (DEVD-CHO) was tested. Cytofluorometric analyses indicate that neither caspase inhibitor had a significant effect on apoptosis (Table IVGo). As control, the activity of ZVAD.fmk was checked by measuring apoptosis of murine thymocytes treated with 10-7 M dexamethasone for 24 h. ZVAD.fmk completely blocked dexamethasone-induced thymocyte apoptosis (Table IVGo). These results imply that caspase-1 and caspase-3 are not involved in the macrophage apoptosis mechanism induced by GBS-III.

Extracellular calcium is involved in GBS induction of apoptosis

Our results suggest that a defect in membrane permeability, caused by GBS ß-hemolysin, could play a role in apoptosis. GBS ß-hemolysin by inducing membrane permeability would allow the influx of ions among which Ca2+. Because growing evidence suggests that the increase in cytosolic Ca2+ level is involved in some apoptosis models (42, 43, 44, 45), we examined the potential role of extracellular Ca2+ influx in GBS-induced macrophage apoptosis using the Ca2+ chelator, EGTA. Incubation of macrophages with EGTA, 1 mM during 2 h infection and 0.5 mM during the following 24 h of culture, resulted in about 70% inhibition of GBS-induced apoptosis, evaluated by cytofluorometric analysis of PI stained DNA (Fig. 9Go). This inhibition of apoptosis by EGTA, can be reversed by an excess of CaCl2 but not MgCl2 during incubation with EGTA (Fig. 9Go).



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  		FIGURE 9. Inhibition of GBS-induced apoptosis by a calcium chelator. Macrophages infected with GBS-III in the absence or presence of 1 mM EGTA, and maintained at 0.5 mM during the experiments. Some cultures were added with 1 mM CaCl2 or MgCl2. At 24 h after infection, apoptosis was measured evaluating the percentage of hypodiploid nuclei at flow cytometry. Treatment of control macrophages with EGTA, CaCl2, MgCl2, and EGTA plus CaCl2 or MgCl2, for 24 h did not induce apoptosis and did not result in loss of cell viability. Data are means ± SD of six experiments performed in triplicate. *, p < 0.01 (GBS-III-infected macrophages treated with EGTA, CaCl2, MgCl2, EGTA plus MgCl2 and EGTA plus CaCl2 vs untreated GBS-III-infected macrophages) according to Student’s t test.

 
PKC is involved in GBS-III-induced macrophage apoptosis

Because PKC activity has a role in modulating apoptosis (46, 47, 48, 49), we determined whether PKC was involved in GBS-induced apoptosis. For this purpose the effect of the potent and selective inhibitors of PKC, GF109203X and calphostin C, on GBS-III-induced macrophage apoptosis was tested. Macrophages were pretreated for 30 min with 1 µM GF109203X or 0.5 µM calphostin C, then infected with GBS-III. After 24 h, the percentage of apoptotic cells was measured by cytofluorometry. Both PKC inhibitors not only failed to inhibit GBS-III-induced apoptosis but enhanced the apoptotic effect of GBS in macrophages. In fact, at 24 h, the percentage of apoptotic cells after PKC inhibition reached about 95% (Fig. 10GoA).



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  		FIGURE 10. Effect of PKC inhibitors and PKC activator on GBS-induced apoptosis. A, Macrophages pretreated for 30 min in the absence or presence of calphostin C (0.5 µM) or GF109203X (1 µM) infected with GBS-III and recovered at 24 h after infection. Apoptosis was determined measuring the percentage of hypodiploid nuclei at flow cytometry. Treatment of control macrophages with inhibitors did not induce apoptosis and did not result in loss of cell viability. Data are means ± SD of six experiments performed in triplicate. B, Macrophages infected with GBS-III in the absence or presence of PMA (1 µg/ml), macrophages pretreated for 30 min with calphostin C (0.5 µM) or GF109203X (1 µM) and infected with GBS-III in the presence of PMA (1 µg/ml), recovered at 24 h after infection. Apoptosis was determined measuring the percentage of hypodiploid nuclei at flow cytometry. Treatment of control macrophages with PMA or PMA plus inhibitors for the same time did not induce apoptosis. Data are means ± SD of six experiments performed in triplicate. *, p < 0.01 (GBS-III-infected macrophages treated with PMA or inhibitors plus PMA vs untreated GBS-III-infected macrophages) according to Student’s t test.

 
Previous studies have demonstrated that PKC activation protects from Ca2+-induced endonuclease activation (42, 45, 50, 51). Because this suggests that the balance between intracellular Ca2+ levels and PKC activation could affect the fate of the cell, the role of PKC activation in GBS-III-induced macrophage apoptosis was analyzed. Macrophages were infected with GBS-III in the presence of PMA (1 µg/ml), an agent known to stimulate PKC, and the percentage of apoptotic cells was measured after 24 h by cytofluorometry. PMA treatment resulted in about 50% inhibition of GBS-III-induced apoptosis (Fig. 10GoB).

To evaluate whether PMA suppression of GBS-induced macrophage apoptosis was directly related to PKC activation the effect of PKC inhibitors, GF109203X and calphostin C, on PMA apoptosis suppression was evaluated. In the presence of GF109203X or calphostin C, the suppressive activity of PMA on macrophage apoptosis by GBS was almost completely abolished (Fig. 10GoB). These results indicate that PKC activation contributes to antagonize the effect of Ca2+ in GBS-induced macrophage apoptosis.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
This study demonstrates that GBS induces apoptosis in murine macrophages. In particular, GBS-III first causes a defect in plasma membrane permeability and, then induces apoptosis, which seems paradoxical because loss of membrane integrity normally leads to immediate death by necrosis. We distinguished apoptosis from necrosis by electron microscopy, gel agarose electrophoresis of fragmented DNA, PI labeling of nuclei and TUNEL. GBS-III-infected macrophages showed the salient features of cells undergoing apoptosis, such as condensation of chromatin and nuclei, cytoplasmic vacuolization, maintenance of organelle structure, DNA fragmentation and loss of DNA stainability. Several different strains, GBS type Ia strain 090, GBS type VI strain 118754 and GBS type III strain COH 1, like GBS-III strain COH31 r/s, also induced macrophage apoptosis indicating that the ability of GBS to cause apoptosis is a common feature to all GBS strains.

GBS also induces apoptosis in human monocytes in the same way as in murine macrophages but does not cause apoptosis in all tumor cell lines analyzed, so indicating that apoptosis induction by GBS is not a general mechanism but is specific for monocytes/ macrophages.

Recent studies have shown that caspases (33, 34, 35) are among the major mediators of apoptosis. Therefore, we tested the effects of two inhibitors, specific for caspase-1 and caspase-3. Results indicate that neither inhibitor affected GBS-induced macrophage apoptosis, suggesting that GBS may trigger apoptosis through an independent caspase-1 and caspase-3 pathway. Other reports have demonstrated caspase-independent apoptosis (19, 34, 52, 53, 54), and proposed that unlike caspase-dependent apoptosis, the signal for triggering apoptosis in these models is integrated within the cells. However, because there are various caspases, other caspase pathways could be involved in GBS-induced apoptosis.

The possibility that GBS-induced apoptosis is mediated by caspase-independent pathways cannot be excluded because it has been reported that some pathways which lead to apoptosis can be activated in a caspase-independent manner. For example, Daxx-ASK connection provides a caspase-independent mechanism for JNK activation by Fas and other stimuli (55, 56, 57) and calpains, calcium dependent proteases, promote apoptosis without activating caspases (58, 59, 60). Therefore, in our model other pathways such as Daxx/ASK/JNK or calpains could be involved in caspase-independent GBS induction of macrophage apoptosis.

The mechanism and strategy by which GBS triggers apoptosis also seems to differ from apoptosis induced by microorganisms such as Shigella and Salmonella. Cell death caused by these pathogens is closely correlated with the ability of the microorganisms to invade infected cells, implying that intracellular localization and the virulence factors mediating cell invasion are also responsible for apoptosis (13, 15, 17). To promote apoptosis GBS does not need to be within the cytoplasm because Cyt D, a drug that prevents bacterial internalization does not affect apoptosis. It would seem that GBS triggers apoptosis from the cell surface. It is possible that GBS either activates the macrophage intrinsic death program or interferes with factors that inhibit the apoptosis program. Alternatively, GBS may produce and/or translocate a factor(s) that induces biochemical changes in the macrophages and triggers apoptosis. In several experimental systems the influx of ions such as Ca2+ has been implicated in the initiation of apoptosis (32, 42, 43, 44, 45). Consistent with the latter hypothesis, GBS-induced apoptosis was not observed when the extracellular Ca2+ was chelated by EGTA or in conditions where plasma membrane permeability defects did not occur, e.g., when we used nonhemolytic gGBS or DPPC, a phospholipid inhibitor of GBS ß-hemolytic activity (10, 36). These results suggest that by inducing membrane alterations GBS could allow an influx of extracellular Ca2+, which triggers apoptosis.

It is well known that GBS has a potent ß-hemolysin strictly bound to the cell surface, which, for its production, requires metabolic activity (36, 41). ß-hemolysin is unstable when released in culture supernatants in the absence of a carrier molecule (41), and, as recently demonstrated, is active against the membrane of some eukaryotic cells (9, 10, 11). In our model GBS ß-hemolysin, like pore-forming proteins (12, 22, 23), perforins (61, 62, 63), and ionophores (43, 47, 49), could be responsible for generating small pores and in these conditions the macrophage membrane would allow the influx of Ca2+, which could directly stimulate Ca2+-dependent endonuclease and initiate apoptosis (12, 31, 32, 42, 43, 44, 45). Because GBS-induced macrophage apoptosis requires host cell protein synthesis, newly synthesized macromolecules are necessary for the transduction of the apoptotic signal. This is in agreement with the postulated mechanism of GBS apoptosis because some of the newly synthesized macromolecules might be endonucleases (29). We also observed a close correlation between the percentage of macrophage apoptosis and hemolytic activity levels (10), of different strains of the microorganism. This further suggests that apoptosis could be a consequence of a membrane permeability defect caused by GBS bound ß-hemolysin.

It is known that pore-forming proteins, perforins, and ionophores contribute to apoptosis induction by causing marked alterations in calcium levels (12, 43, 47, 49, 61, 62, 63), but also by affecting directly or indirectly cellular metabolic processes, ATP levels and mitochondrial function (64, 65, 66, 67). We do not know whether GBS ß-hemolysin has any of these effects and there are no reports in the literature to account for these findings. Therefore, further studies are necessary to understand whether GBS ß-hemolysin, like pore-forming proteins and ionophores could contribute to apoptosis also affecting ATP levels, mitochondrial function, and metabolic processes.

The observation that metabolic inhibition by sodium merthiolate and sodium fluoride abolished the ability of GBS to cause plasma membrane permeability defects and apoptosis (data not shown) indicates that apoptosis induction requires that GBS is metabolically active. However, because sodium merthiolate and sodium fluoride also cause a strong reduction of hemolytic activity (data not shown), our data confirm that continued metabolic activity is necessary both for GBS hemolytic activity expression, as reported also by Marchlewicz and Ducan (36), and for induction of apoptosis.

There is increasing evidence that modulation of PKC activity by several agents affects apoptosis induction. It has also been demonstrated in several cell models that PKC activation protects cells from Ca2+-induced endonuclease activation (31, 42, 45, 50, 51). There is conflicting evidence about PKC involvement in protecting from apoptosis or inducing the apoptotic process (46, 47, 48, 49, 50, 51). In view of our previous results, demonstrating that GBS deactivated the PKC-dependent signal transduction pathway (8), and reports by other authors indicating that the PKC status may play a role in inducing calcium-dependent apoptosis (31, 50, 51), we analyzed the effect of PKC modulators on GBS-induced apoptosis. PKC inhibitors GF109203X and calphostin C failed to inhibit apoptosis and enhanced the apoptotic effect of GBS in macrophages, whereas treatment with PMA, a PKC activator, partially prevented induction of macrophage apoptosis by GBS. These findings suggest that the balance between PKC activation and intracellular Ca2+ concentration is a crucial factor for GBS apoptosis induction.

In conclusion, this study provides the first evidence that GBS induces apoptosis in immune cells, and emphasizes the complexity of the strategy used by GBS to overcome host immune defenses. Until now, the most known GBS pathogenic mechanism was avoiding phagocytosis by an anti-phagocytic capsule (1, 2, 3). Recently, it has been demonstrated that GBS can survive in different cell types (4, 5, 6, 7, 8) and kill endothelial and epithelial cells by necrosis (9, 10, 11). This study demonstrates that GBS can also induce an intrinsic cell death program in macrophages. The ability of GBS to kill macrophages and human monocytes by apoptosis could be an important pathogenic mechanism by which the microorganism evades host immune defenses and causes disease.


   Footnotes
 
1 This work was supported in part by Perugia Ateneo Funds (Program for Young Researchers, 1999) and Ministero dell’Università e della Ricerca Scientifica e Tecnologica (Funds ex 40%, 1997), Italy. Back

2 Address correspondence and reprint requests to Prof. Pierfrancesco Marconi, Department of Clinical Medicine, Pathology and Pharmacology, General Pathology and Immunology Section, University of Perugia, General Hospital, Monteluce, 06100 Perugia, Italy. Back

3 Abbreviations used in this paper: GBS, Group B Streptococcus; PKC, protein kinase C; Ca2+, calcium ions; DPPC, dipalmitoylphosphatidylcholine; PI, propidium iodide; CHX, cycloheximide; Cyt D, cytochalasin D; Act D, actinomycin D; ZVAD.fmk, caspase 1-like protease inhibitor V; DEVD-CHO, inhibitor of CPP32/apopain caspase 3; GBS-III, GBS type III strain COH31 r/s; gGBS-III, GBS-III grown in the presence of glucose; THB, Todd-Hewitt broth. Back

Received for publication July 29, 1999. Accepted for publication July 3, 2000.


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