Depletion of Alveolar Macrophages Exerts Protective Effects in Pulmonary Tuberculosis in Mice

Jaklien C. Leemans, Nicole P. Juffermans, Sandrine Florquin, Nico van Rooijen, Margriet J. Vervoordeldonk, Annelies Verbon, Sander J. H. van Deventer and Tom van der Poll
J Immunol April 1, 2001, 166 (7) 4604-4611; DOI: https://doi.org/10.4049/jimmunol.166.7.4604

Abstract

Mycobacterium tuberculosis bacilli are intracellular organisms that reside in phagosomes of alveolar macrophages (AMs). To determine the in vivo role of AM depletion in host defense against M. tuberculosis infection, mice with pulmonary tuberculosis induced by intranasal administration of virulent M. tuberculosis were treated intranasally with either liposome-encapsulated dichloromethylene diphosphonate (AM− mice), liposomes, or saline (AM+ mice). AM− mice were completely protected against lethality, which was associated with a reduced outgrowth of mycobacteria in lungs and liver, and a polarized production of type 1 cytokines in lung tissue, and by splenocytes stimulated ex vivo. AM− mice displayed deficient granuloma formation, but were more capable of attraction and activation of T cells into the lung and had increased numbers of pulmonary polymorphonuclear cells. These data demonstrate that depletion of AMs is protective during pulmonary tuberculosis.

Mycobacterium tuberculosis is responsible for much morbidity and mortality worldwide (1). The increasing incidence of antibiotic resistance, together with synergism between HIV and tuberculosis, has heightened our interest in this important infectious disease and in mechanisms contributing to antimicrobial host defense.

Mycobacteria are intracellular pathogens that are taken up by host alveolar macrophages (AMs),3 in which they either are killed or survive. Surviving bacilli start to proliferate and are released, leading to infection of additional host cells. Apoptosis of AMs could be an effective weapon to kill or inhibit the growth of intracellular mycobacteria. Several findings suggest that AM apoptosis plays an important role in tuberculosis. Infection of human AMs with M. tuberculosis has been shown to induce apoptosis in vitro (2). Furthermore, extensive apoptosis (50–70%) was found within tuberculous granulomas in lungs of tuberculosis patients (2), and a significant increase in the number of apoptotic AMs was observed in bronchoalveolar lavage fluid (BALF) from patients with active pulmonary tuberculosis (3, 4). Despite these observations, it is not clear which role AM apoptosis plays in the pathobiology of this disease and whether it increases or decreases the mycobacterial load in vivo. In vitro studies suggest that apoptosis may be a macrophage defense mechanism to infection by mycobacteria. Indeed, apoptosis of human monocytes limited the growth of Mycobacterium avium (5), Mycobacterium bovis bacillus Calmette-Guérin (6), and M. tuberculosis (7). However, in vitro studies are not adequate to determine the net effect of AM depletion on the host response to tuberculosis. AMs have important phagocytic and immune functions that could be disturbed by the apoptotic process. Clearance of microorganisms that reach the alveolar space relies partly on phagocytic AMs. Furthermore, macrophages present mycobacterial Ags to CD4+ T lymphocytes that are central in the acquired resistance to M. tuberculosis. Macrophages are a significant source of type 1 cytokines during mycobacterial infection (8), which are known to be important for the development of protective immunity (9). In addition, AMs produce IFN-γ in response to M. tuberculosis (10), which is a pivotal mediator in host resistance to tuberculosis (11, 12). Finally, mononuclear cells are involved in the formation of granulomas, which are critical in restricting mycobacterial growth and dissemination (13). Hence, theoretically AM depletion could have beneficial and detrimental effects during tuberculosis in vivo.

In the present study, we determined the role of AM depletion in M. tuberculosis infection in mice, using the well-validated method of intrapulmonary delivery of liposome-encapsulated dichloromethylene diphosphonate (clodronic-acid disodium salt tetrahydrate, CL2MBP). Intratracheal administration of liposome-encapsulated CL2MBP selectively depletes AMs (14) by apoptosis (15, 16) without damaging other cell types in the lung (17). In this work, we present the first evidence that AM depletion in vivo leads to improved clearance of M. tuberculosis bacilli.

Materials and Methods

Mice

Pathogen-free 6-wk-old female BALB/c mice were obtained from Harlan Sprague-Dawley (Horst, The Netherlands) and were maintained in biosafety level 3 facilities. The Animal Care and Use Committee of the University of Amsterdam (Amsterdam, The Netherlands) approved all experiments.

Experimental infection

A virulent laboratory strain of M. tuberculosis H37Rv was grown in liquid Dubos medium containing 0.01% Tween 80 for 4 days. A replicate culture was incubated at 37°C, harvested at mid-log phase, and stored in aliquots at −70°C. For each experiment, a vial was thawed and washed twice with sterile 0.9% NaCl. Mice were anesthetized by inhalation with isoflurane (Abbott Laboratories, Kent, U.K.) and infected with 1 × 105 live bacilli in 50 μl saline, as determined by viable counts on 7H11 Middlebrook agar plates. Bacterial administration was performed intranasally (i.n.), as described previously (18, 19, 20). Groups of eight mice per time point were sacrificed 2 or 5 wk postinfection, and lungs and one lobus of the liver were removed aseptically. Organs were homogenized with a tissue homogenizer (Biospec Products, Bartlesville, OK) in 5 vol of sterile 0.9% NaCl, and 10-fold serial dilutions were plated on Middlebrook 7H11 agar plates to determine bacterial loads. Colonies were counted after 21-day incubation at 37°C. Numbers of CFUs are provided as total in the lungs or as total per gram liver. For cytokine measurements, lung homogenates were diluted 1/1 in lysis buffer (150 mM NaCl, 15 mM Tris, 1 mM MgCl.H2O, 1 mM CaCl2, 1% Triton X-100, 100 μg/ml pepstatin A, leupeptin, and aprotinin), and incubated on ice for 30 min. Supernatants were sterilized using a 0.22-μm filter (Corning, Corning, NY) and frozen at −20°C until assays were performed.

In vivo AM depletion

CL2MBP was a gift from Roche Diagnostics (Mannheim, Germany). Preparation of liposomes containing CL2MBP was done as described previously (17). For assessment of AM depletion, five uninfected mice per group were i.n. inoculated with 100 μl of 0.9% NaCl, PBS liposomes, or CL2MBP liposomes. Two days later, AMs were quantified in the BALF. For the tuberculosis experiments, 100 μl saline, PBS liposomes, or CL2MBP liposomes were instilled 2 days before and 6, 14, and 25 days after M. tuberculosis challenge.

Detection of apoptotic cells

To confirm apoptotic cell death induced by CL2MBP liposomes, a cleavage of poly(ADP-ribose) polymerase (PARP) was determined, as described previously (21). Briefly, 7 h after i.n. instillation of CL2MBP liposomes, tissue Tek OTC compound (Miles Scientific, Naperville, IL) was instilled intratracheally into lungs, which were then snap frozen and stored at −70°C. Cryostat sections (7 μm) of frozen lungs were fixed in cold acetone for 10 min, incubated with 0.3% H2O2 in methanol for 15 min, blocked for nonspecific Ig binding by incubation for 30 min with a 1/10 dilution of normal goat serum, and incubated overnight with rabbit anti-PARP cleavage site (214/215)-specific Ab (Biosource International, Camarillo, CA; 5 μg/ml). This was followed by a 30-min incubation with poly-HRP goat anti-rabbit IgG (Immunovision, Springdale, AZ). The peroxidase activity was revealed by adding AEC substrate (3-amino-9-ethyl-carbazole; Sigma, Buchs, Switzerland) and H2O2. Sections were counterstained with hematoxylin. Negative controls were established by adding nonspecific isotype controls as primary Abs.

Assessment of in vitro effect of CL2MDP liposomes on M. tuberculosis

A total of 2.5 × 103 CFUs was incubated in octuplicate in 96-well round-bottom culture plates in the presence of Lowenstein-Jensen medium (Becton Dickinson, Franklin Lakes, NJ) with either 0.9% NaCl, PBS liposomes, or CL2MBP liposomes. After 48-h incubation at 37°C in 5% CO2, colonies were counted.

Lung lavage

Bronchoalveolar lavage was performed to obtain intraalveolar cells. Briefly, mice were anesthetized, and the trachea was exposed through a midline incision and cannulated with a sterile 22-gauge Abbocath-T catheter (Abbott, Sligo, Ireland). The lungs were then lavaged with two 0.5-ml aliquots of sterile 0.9% NaCl. A total of 0.9–1 ml of lavage fluid was retrieved per mouse, and total leukocyte count was determined using a hemacytometer and TÜRK’s solution (Merck, Gibbstown, NJ). BALFs from infected mice were fixed with 2% paraformaldehyde. The number of AMs, polymorphonuclear cells (PMNs), and lymphocytes were calculated from these totals, using cytospin preparations stained with modified Giemsa stain (Diff-Quick; Baxter, McGaw Park, IL).

Histological analyses

The left lungs were removed 2 or 5 wk after inoculation with M. tuberculosis and fixed in 4% paraformaldehyde in PBS for 24 h. One lobus of the liver of noninfected mice was removed 2 days after CL2MBP liposome treatment. After embedding in paraffin, 4-μm-thick sections were stained with eosin hematoxylin-eosin or the Ziehl-Neelsen (ZN) stain for acid fast bacilli. All slides were coded and semiquantitatively scored for the total area of inflammation (percentage of surface of the slide) and granuloma format by a pathologist.

FACS analysis

Lung cells from mice 2 and 5 wk postinfection (eight mice per group) were analyzed by FACS (Becton Dickinson). Pulmonary cell suspension was obtained using an automated disaggregation device (Medimachine System; Dako, Glostrup, Denmark) and resuspended in medium (RPMI 1640 (BioWhittaker, Belgium), 10% FCS, 1% antibiotic-antimycotic (Life Technologies, Rockville, MD)). Cells from two mice per group were pooled for each time point (yielding four samples for FACS analysis per group) and were brought to a concentration of 4 × 106 cells/ml FACS buffer (PBS supplement with 0.5% BSA, 0.01% NaN3, and 100 mM EDTA). Immunostaining for cell surface molecules was performed for 30 min at 4°C using directly labeled Abs against CD3 (anti-CD3 PE), CD4 (anti-CD4 CyChrome), CD8 (anti-CD8 FITC, anti-CD8 PerCP), CD25 (anti-CD25 FITC), CD69 (anti-CD69 FITC), and Gr-1 (anti-Gr-1 FITC). All Abs were used in concentrations recommended by the manufacturer (PharMingen, San Diego, CA). To correct for aspecific staining, an appropriate control Ab (rat IgG2; PharMingen) was used. Cells were fixed with 2% paraformaldehyde, T cells were analyzed by gating the CD3+ population, and granulocytes by gating the forward and side angle scatter-gated PMN population. The number of positive cells was obtained by setting a quadrant marker for nonspecific staining.

Splenocyte stimulation

Single cell suspensions were obtained by crushing spleens through a 40-μm cell strainer (Becton Dickinson). Erythrocytes were lysed with ice-cold isotonic NH4Cl solution (155 mM NH4Cl, 10 mM KHCO3, 100 mM EDTA, pH 7.4), and the remaining cells were washed twice. Splenocytes were suspended in medium (RPMI 1640 (BioWhittaker), 10% FCS, 1% antibiotic-antimycotic (Life Technologies)), seeded in 96-well round-bottom culture plates at a cell density of 5 × 105 cells in triplicate, and stimulated with 20 μg/ml tuberculin-purified protein derivative (PPD; Statens Seruminstitut, Copenhagen, Denmark). Supernatants were harvested after a 48-h incubation at 37°C in 5% CO2, and cytokine levels were analyzed by ELISA.

Splenocyte proliferation assay

Proliferation of splenocytes was measured by the MTT assay, which measures reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide to formasan in mitochondria of viable cells (22). Splenocytes were seeded in triplicate at a density of 5 × 105 cells/well in flat-bottom 96-well plates and stimulated with 20 μg/ml PPD. After 42 h at 37°C in 5% CO2, cells were incubated with 5 mg/ml MTT (Sigma, St. Louis, MO) in PBS (pH 7.2) for an additional 6 h. Supernatants were decanted, and the formazan precipitates were solubilized by the addition of 0.04 N HCl in isopropanol and placed on a plate shaker for 10 min, after which cells were dissolved in 2% paraformaldehyde. Cell proliferation was quantified using an ELISA reader at 570 nm. The absorbance of the untreated cultures was set at 100%.

Cytokine measurements

Cytokines were measured in lung homogenates and spleen cell supernatants by specific ELISAs using matched Ab pairs according to the manufacturer’s instructions: IFN-γ, IL-2, IL-4 (R&D Systems, Minneapolis, MN), and IL-10 (PharMingen).

Statistical analysis

All values are expressed as mean ± SEM. Comparisons were done with Mann-Whitney U tests. For comparison of survival curves, Kaplan-Meier analysis with a log rank test was used. Values of p ≤ 0.05 were considered statistically significant.

Results

Effects of AM depletion on the course of infection

Intranasal administration of liposome-encapsulated CL2MBP resulted in >70% AM depletion in BALF of uninfected mice after 2 days (Fig. 1⇓a). Liposome treatment showed no effect on macrophage numbers. Lungs of CL2MBP liposome-treated mice presented large areas of degenerated macrophages with cell debris and apoptotic bodies in the alveolar spaces. This result is in line with previous reports on the capacity of intratracheally administered CL2MBP liposomes or CL2MBP liposomes given by aerosol to deplete AMs (23, 24). Induction of AM apoptosis by CL2MBP liposome treatment was confirmed by the detection of cleaved PARP in lung tissue (Fig. 1⇓b).

FIGURE 1.
FIGURE 1.

a, Effect of CL2MBP liposomes on AMs 2 days after i.n. administration. The bars indicate the percentage of total AMs remaining in AM− mice (▪) or AM+ (liposomes) mice (▨) compared to mice receiving saline (□; 100%). The data are presented as mean and SEM from five mice per group. ∗, p < 0.05 AM− mice vs AM+ (saline) mice; †, p < 0.05 AM− mice vs AM+ (liposomes) mice. b, Apoptotic AMs were visualized by immunostaining lung tissue for cleaved PARP 7 h after CL2MBP liposome treatment (original magnification, ×80).

To investigate the role of AMs in the outcome of tuberculosis, CL2MBP liposomes were given i.n. 2 days before, and 6, 14, and 25 days after induction of tuberculosis (AM− mice). Mice given saline or liposomes served as controls (AM+ mice). Survival and bacterial load in lungs and liver were analyzed to determine resistance to tuberculosis. As shown in Fig. 2⇓, survival in AM+ (saline) mice decreased extensively from day 35 onward, resulting in 90% mortality after 5 mo. In sharp contrast, all AM− mice controlled the same infectious dose and survived the 5-mo follow-up (p < 0.0001). In addition, a significant decrease in survival was found in the AM+ (liposomes) mice compared with AM− animals (p = 0.029).

FIGURE 2.
FIGURE 2.

Effect of AM depletion on survival of mice following M. tuberculosis infection. BALB/c mice (n = 10 per group) were i.n. administered with saline, liposomes, or CL2MBP liposomes prior to and after bacterial challenge of 1 × 105 M. tuberculosis H37Rv. ∗, p < 0.05 AM− mice vs AM+ (saline) mice; †, p < 0.05 AM− mice vs AM+ (liposomes) mice; ‡, p < AM+ (saline) mice vs AM+ (liposomes) mice.

Because of the impressive differences in survival, we determined whether differences existed in mycobacterial load during earlier phases of the infection. The number of bacteria deposited in the lungs and liver was determined 1 day after infection. Bacterial counts in the lungs were comparable with the numbers of bacteria that were given intranasally and did not differ between the groups. The numbers of M. tuberculosis CFUs recovered from lungs were not significantly different between AM+ (saline/liposomes) and AM− mice 2 wk postinfection (Fig. 3⇓a). At 5 wk postinfection, significant differences in tissue content of M. tuberculosis bacilli were observed between AM+ and AM− mice. The lungs of AM− mice contained 9.7-fold less viable mycobacteria than those of AM+ (saline) animals (p = 0.021, Fig. 3⇓a) and 7.1-fold less than of AM+ (liposomes) mice (p = 0.035).

FIGURE 3.
FIGURE 3.

Growth of M. tuberculosis in lungs (a) and liver (b) of AM+ (saline/liposomes) mice and AM− mice. Data are mean and SEM of CFUs from eight mice per group for each time point. ∗, p < 0.05 AM− mice vs AM+ (saline) mice; †, p < 0.05 AM− mice vs AM+ (liposomes) mice.

To give more clarity on the residing place of mycobacteria in AM− animals, lavage fluids of these mice were analyzed with a Ziehl-Neelsen (ZN) stain for acid fast bacilli. In BALFs of AM+ mice, mycobacteria were present within the cytoplasm of 39 ± 4% (2 wk postinfection) and 47 ± 10% (5 wk postinfection) of the macrophages and in few PMNs. In the BALFs of AM− mice, we found extracellular mycobacteria, but mycobacteria were especially present in cell debris.

At 2 wk postinfection, the mycobacterial load in the liver of AM− mice was increased in comparison with AM+ (saline) mice (p = 0.02) and AM+ (liposomes) mice (p = 0.004, Fig. 3⇑b). At 5 wk postinfection, the number of organisms in the liver of AM− mice was 3.6 times lower than that in AM+ (saline) mice (p = 0.011, Fig. 3⇑b) and 2.7 times lower than in AM+ (liposomes) animals (not significantly different). Bacterial counts in lungs and liver of the control groups treated with either saline or liposomes mice were not significantly different at either time point.

To exclude the possibility that liposome-encapsulated CL2MBP had a direct effect on mycobacteria, M. tuberculosis was incubated in vitro in the presence or absence of this agent for 2 days. Bacterial counting demonstrated no direct antimycobacterial effect of liposome-encapsulated CL2MBP (data not shown).

Together these findings suggest that AM depletion by apoptosis can play an important role in controlling M. tuberculosis infection.

Histology

Two weeks after M. tuberculosis inoculation, lungs of AM+ (saline/lipsomes) mice exhibited more or less well-defined granulomas comprising a majority of epithelioid and foamy cells and a small number of lymphocytes throughout the parenchyma (Fig. 4⇓a). Dense lymphocytic infiltrates were also present around small vessels. Lungs of AM− mice showed a relatively diffuse infiltrate of granulocytes with prominent perivascular lymphocytic infiltrates. Well-defined granulomas were not present (Fig. 4⇓b). The percentage of inflamed parenchyma was similar in all groups (AM+ (saline) mice, 21.25 ± 3.5%; AM+ (liposomes) mice, 16.9 ± 3.5%; AM− mice, 18.8 ± 2.9%).

FIGURE 4.
FIGURE 4.

Lung histopathology. a, Two weeks postinfection, lungs of AM+ mice showed well-shaped granulomas, chiefly composed by macrophages and few lymphocytes (hematoxylin and eosin staining; original magnification, ×50). b, At the same time point, lungs of AM− mice displayed slight perivascular lymphocytic infiltrates and degenerated macrophages in alveolar spaces. Well-defined granulomas were not found (hematoxylin and eosin staining; original magnification, ×50). c, Five weeks postinfection, lungs of AM+ mice displayed a diffuse inflammatory infiltrate predominantly composed of macrophages (hematoxylin and eosin staining; original magnification, ×25). d, Lungs of AM− mice presented an almost normal aspect (hematoxylin and eosin staining; original magnification, ×25). The slides shown are representative for a total of eight mice per group for each time point.

After 5 wk, the inflammatory infiltrates in lungs of all mice became more diffuse and intense with a cellular composition comparable in AM+ (saline/liposomes) and AM− mice. However, the percentage of inflamed parenchyma was less in AM− mice (45 ± 4.8%) than in AM+ mice (saline, 61.9 ± 4.9%; liposomes, 62.9 ± 7.1%) (Fig. 4⇑, c and d).

Cell subsets

CD4+ T cells have an established role in protective immunity against M. tuberculosis infection (25, 26, 27), and must be stimulated with specific ligands on the surface of APCs. To study whether AMs are important for the induction of CD4+ T cell-mediated immunity, we investigated the phenotypes of immune cells in total lungs by FACS analysis. As shown in Table I⇓, the percentages of CD4+ T cells did not differ between AM+ and AM− mice 5 wk postinfection and were slightly reduced in AM− mice 2 wk postinfection. Two weeks after infection, CD4+ lymphocytes of AM− mice were demonstrated to be more activated than CD4+ lymphocytes of AM+ mice, as assessed by the activation markers CD69 and CD25.

View this table:
Table I.

Effect of AM depletion on cell subsets in total lungs during tuberculosisa

Besides CD4+ T cells, CD8+ T cells have also been suggested to participate in host defense against mycobacterial infections (25). The percentage of CD8+ T cells in lung homogenates was not changed in AM− mice as compared with AM+ (saline/liposomes) mice 5 wk postinfection and slightly increased 2 wk postinfection. The expression of the activation marker CD69 on these cells was slightly increased. CD25 expression on CD8+ cells could not be detected. The absolute number of PMNs was higher in AM− mice than in AM+ control animals 2 wk postinfection. At 5 wk, numbers of total leukocytes were decreased in AM− mice compared with AM+ (saline/liposomes) mice, probably reflecting disease severity.

To obtain more insight into the leukocyte influx into the alveolar compartment, lungs were lavaged, cells were counted, and cytospin preparations were stained with eosin hematoxylin-eosin (Table II⇓). Two weeks postinfection, the number of leukocytes was higher in AM− mice than in AM+ (saline/liposomes) mice. In line with the numbers of leukocytes in total lungs at 5 wk postinfection, cell numbers in BALFs were lower in AM− mice than in AM+ (saline/liposomes) mice. As could be expected, CL2MBP liposome-treated animals had 2 times less AMs in their BALFs than AM+ (saline/liposomes) mice. The amount of PMNs and lymphocytes in AM− mice was however increased 2 wk postinfection in comparison with AM+ (saline/liposomes) mice. As a consequence of lower leukocyte numbers in AM− mice 5 wk postinfection, numbers of PMNs and lymphocytes were decreased in this group compared with AM+ (saline/liposomes) mice.

View this table:
Table II.

Effect of AM depletion on cellular composition on BALFs during tuberculosisa

Cytokine expression patterns in lung

Since development of early Th1 cellular immunity is essential for the elimination of M. tuberculosis (9), we investigated whether the improved outcome of tuberculosis seen in the AM− mice was associated with a shift in cytokine production early in the infection. We therefore measured the concentrations of Th1 (IFN-γ and IL-2) and Th2 (IL-4, IL-10) cytokines in the lung. As shown in Fig. 5⇓, all cytokines were reduced in AM− mice compared with AM+ (saline/liposomes) mice 2 wk postinfection. Importantly, when compared with AM+ (saline/liposomes) mice, Th2 cytokine concentrations were relatively more reduced than the levels of Th1 cytokines in AM− mice. As a consequence, a more profound Th1 response was found in lungs of AM− mice.

FIGURE 5.
FIGURE 5.

The effect of AM depletion on M. tuberculosis-mediated induction of type 1 cytokines (a) and type 2 cytokines (b) in lungs of AM+ (saline) mice (□), AM+ (liposomes) mice (▨), and AM− mice (▪) 2 wk postinfection. The data represent the mean and SEM of eight mice. ∗, p < 0.05 AM− mice vs AM+ (saline) mice; †, p < 0.05 AM− mice vs AM+ (liposomes) mice.

Cytokine and proliferative response of ex vivo stimulated spleen cells

The ability of spleen cells, harvested 2 wk postinfection with M. tuberculosis to produce cytokines ex vivo upon stimulation with PPD, was investigated as another measure of Th1 vs Th2 response. Spleen cells from AM− mice secreted 3.5-fold higher levels of IFN-γ than splenocytes from AM+ (saline) mice and 2-fold higher levels than splenocytes from AM+ (liposomes) mice (Fig. 6⇓). IL-4 was not detectable in supernatants of PPD-stimulated splenocytes in all groups. When stimulated with coated anti-CD3 and anti-CD28 Abs, splenocytes from AM− mice secreted higher levels of IFN-γ and significantly lower levels of IL-4 compared with AM+ (saline) animals. In addition, the proliferation responses of splenocytes to PPD were estimated using the MTT incorporation assay. We found that splenocytes from AM− mice induced the strongest proliferative response to PPD, although not statistically significant (AM+ (saline) mice, 155 ± 17%; AM+ (liposomes) mice, 158 ± 6%; and AM− mice, 195 ± 31%).

FIGURE 6.
FIGURE 6.

Splenocytes from infected AM− mice (▪) release more IFN-γ in response to PPD and anti-CD3/28 Abs and less IL-4 in response to anti-CD3/28 ABS than splenocytes from infected AM+ mice (□, saline; ▨, liposomes). Splenocytes were harvested 2 wk after i.n. inoculation with M. tuberculosis, and stimulated for 48 h. The data are mean and SEM of eight mice per group. ∗, p < 0.05 AM− mice vs AM+ (saline) mice; †, p < 0.05 AM− mice vs AM+ (liposomes) mice; ‡, p < AM+ (saline) mice vs AM+ (liposomes) mice.

Discussion

AMs may have a dual role during infection with M. tuberculosis. On the one hand, AMs have several tools to combat intracellular pathogens, such as the production of IFN-γ, and toxic effector molecules (reactive oxygen intermediates and reactive nitrogen intermediates), and the deprivation of the intracellular iron availability. On the other hand, mycobacteria may in part rely on the intracellular environment of AMs for their multiplication. We demonstrate in this study that depletion of AMs in vivo improves the outcome of pulmonary tuberculosis, as indicated by a total protection against lethality and an attenuated outgrowth of mycobacteria in lungs. These results suggest that AMs facilitate the growth of M. tuberculosis in the pulmonary compartment, and that AM apoptosis may be part of the host defense mechanisms during tuberculosis. Interestingly, AMs do seem to have a significant role in the initial capturing of mycobacteria, as indicated by the observation that 2 wk postinfection AM− mice had more mycobacteria in their livers.

Host cell apoptosis has already been demonstrated to be a defense strategy to limit the growth of viruses, which like mycobacteria live intracellularly (28, 29, 30). The fact that AM apoptosis might contribute to host defense is further supported by observations of an inverse relationship between apoptosis and virulence, i.e., the virulent M. tuberculosis strain H37Rv induced less apoptosis upon human AM infection than the attenuated H37Ra strain (2). Hence, mycobacteria seem to have developed ways to modulate the protective apoptotic process of AMs, and pathogen-induced suppression of the host cell-death pathway may serve to evade host defenses that can act to limit the infection. It should be noted that the role of AMs in respiratory infections by extracellularly growing pathogens is opposite. Indeed, induction of AM apoptosis during Klebsiella pneumonia impaired host defense mechanisms (31).

The most straightforward interpretation of the improved tuberculosis outcome in AM− mice is that AM depletion reduces the viability of M. tuberculosis because the environment for intracellular replication and hiding is destroyed (32). Furthermore, apoptotic bodies maintain their plasma membrane integrity so that bacilli are contained from the extracellular environment and can be engulfed by newly recruited AMs (5). A further explanation for the protection observed with AM depletion may be that the early immune response was dominated by a Th1-type profile that is essential for resistance to mycobacteria (9). The predominance of Th1-type cytokines in AM− mice existed both in lung tissue, in which especially the concentrations of Th2 cytokines were decreased, and in supernatants of PPD-stimulated splenocytes. Wang et al. (8) recently reported that lung macrophages harvested during mycobacterial infection release significant amounts of type 1 cytokines. In line with these observations, we found lower levels of IFN-γ and IL-2 in lung homogenates 2 wk postinfection. However, the net in vivo effect of AM depletion was a relative type 1 dominance in the lung. It is unlikely that this type 1 shift was the consequence of a milder inflammatory response in AM− mice, which, if anything, showed slightly more evidence of inflammation (Tables I⇑ and II⇑). It is conceivable that the depletion of AMs was involved in this shift. AMs are typical macrophage populations, which are known to induce differentiation of naive T cells into Th2 type cells, and to exert Th2-associated effector functions (33, 34). It is furthermore reasonable that the reduced IL-10 levels in AM− mice were involved in this shift, considering that IL-10 is derived from AMs (35, 36) and that this cytokine down-regulates type 1 cytokine production (37). Another reason for the better outcome in AM− mice could be the enhanced influx of PMNs and activated lymphocytes into the lung of AM− mice (Tables I⇑ and II⇑). T cells have a prominent role in the protective immunity against M. tuberculosis (38), and therefore increased numbers of these cells most likely contribute to resistance. In support of a role for PMNs in mycobacterial infections, a protective function of a neutrophilic response was demonstrated by increased susceptibility of mice depleted of neutrophils to M. tuberculosis (39). It is conceivable that with the lack of phagocytosing AMs in AM− mice, PMNs get more signals to migrate to the lung to ingest and eliminate the apoptotic bodies derived from AMs.

AM− mice were fully or even more capable of attraction and activation of T cells in the pulmonary compartment. In this context, it should be noted that AMs are poor APCs (40, 41, 42, 43) and that dendritic cells (44, 45) and interstitial macrophages (46) are considered the most efficient APCs in the lung. In vitro experiments even point out that AMs are highly T cell suppressive (47, 48, 49). It is conceivable that the suppressive effects of AMs on the pulmonary immune response may serve to limit damage caused by severe immune responses in lung tissue, but at the same time may impair host defense during tuberculosis.

Two weeks postinfection, more M. tuberculosis CFUs were recovered from livers of AM− mice than from AM+ mice. Our study does not elucidate the mechanisms contributing to this observation. Possibly, bacilli that remain extracellularly (such as in AM− mice) gain access to the blood and lymphatic circulation more easily. It is unlikely that CL2MBP liposome inhalation influenced the number and/or function of Kupffer cells in the liver, considering that liposomes are not able to cross capillary walls and other vascular barriers (17). The absence of an effect of inhaled CL2MBP liposomes on Kupffer cells is supported by the fact that liver histology did not differ between different treatment groups (data not shown).

Liposomes were used to encapsulate Cl2MBP, because these phospholipid spheres are eagerly taken up by macrophages. Liposomes can reduce the phagocytic and migratory behavior of AMs (50) and may therefore influence host defense against M. tuberculosis. In accordance, animals treated i.n. with liposomes only (i.e., without CL2MBP), displayed an enhanced survival and a slight (not significant) reduction in M. tuberculosis CFU in lungs and liver compared with AM+ (saline) mice. Since we sought to determine the role of AMs in pulmonary tuberculosis, control mice should have normal, nonsuppressed AMs (17), and in this way we consider AM+ (saline) animals better controls than AM+ (liposome) mice. Physical depletion of AMs with CL2MBP liposomes ensures that all functions of the macrophages that have ingested this compound are abrogated. AMs that phagocytosed the liposomes alone are expected to have some functional disabilities. Nonetheless, AM+ liposome-treated mice differed significantly from AM− mice with respect to all responses measured.

This study is the first to show that AM depletion in vivo is protective in M. tuberculosis infection and that it is associated with an enhanced Th1-mediated immune response. AM apoptosis as observed in patients with tuberculosis could therefore be an important antimycobacterial defense process. The present results not only provide new insights into possible macrophage antimicrobial defense mechanisms, but also reveal potentially new therapeutic strategies to manage intracellular bacterial diseases.

Acknowledgments

We thank Joost Daalhuisen, Nike Claessen, and Dr. Arend Kolk for expert technical assistance.

Footnotes

  • ↵1 This work was supported by a grant from The Netherlands Organization for Scientific Research (to J.C.L.) and a Fellowship from the Royal Dutch Academy of Arts and Sciences (to T.v.d.P.).

  • ↵2 Address correspondence and reprint requests to Dr. Jaklien Leemans, Laboratory of Experimental Internal Medicine, Academic Medical Center, University of Amsterdam, G2-105, Meibergdreef 9, 1105 AZ Amsterdam, The Netherlands. E-mail address: j.c.leemans{at}amc.uva.nl

  • ↵3 Abbreviations used in this paper: AM, alveolar macrophage; BALF, bronchoalveolar lavage fluid; Cl2MBP, dichloromethylene bisphosphonate; i.n., intranasal; PARP, poly(ADP-ribose) polymerase; PMN, polymorphonuclear cell; PPD, purified protein derivative.

  • Received April 14, 2000.
  • Accepted January 22, 2001.

References

  1. ↵
    Dye, C., S. Scheele, P. Dolin, V. Pathania, M. C. Raviglione. 1999. Consensus statement: global burden of tuberculosis: estimated incidence, prevalence, and mortality by country: WHO Global Surveillance and Monitoring Project. J. Am. Med. Assoc. 282: 677
    OpenUrlCrossRefPubMed
  2. ↵
    Keane, J., M. K. Balcewicz-Sablinska, H. G. Remold, G. L. Chupp, B. B. Meek, M. J. Fenton, H. Kornfeld. 1997. Infection by Mycobacterium tuberculosis promotes human alveolar macrophage apoptosis. Infect. Immun. 65: 298
    OpenUrlAbstract/FREE Full Text
  3. ↵
    Klingler, K., K. M. Tchou-Wong, O. Brandli, C. Aston, R. Kim, C. Chi, W. N. Rom. 1997. Effects of mycobacteria on regulation of apoptosis in mononuclear phagocytes. Infect. Immun. 65: 5272
    OpenUrlAbstract/FREE Full Text
  4. ↵
    Placido, R., G. Mancino, A. Amendola, F. Mariani, S. Vendetti, M. Piacentini, A. Sanduzzi, M. L. Bocchino, M. Zembala, V. Colizzi. 1997. Apoptosis of human monocytes/macrophages in Mycobacterium tuberculosis infection. J. Pathol. 181: 31
    OpenUrlCrossRefPubMed
  5. ↵
    Fratazzi, C., R. D. Arbeit, C. Carini, H. G. Remold. 1997. Programmed cell death of Mycobacterium avium serovar 4-infected human macrophages prevents the mycobacteria from spreading and induces mycobacterial growth inhibition by freshly added, uninfected macrophages. J. Immunol. 158: 4320
    OpenUrlAbstract
  6. ↵
    Molloy, A., P. Laochumroonvorapong, G. Kaplan. 1994. Apoptosis, but not necrosis, of infected monocytes is coupled with killing of intracellular bacillus Calmette-Guérin. J. Exp. Med. 180: 1499
    OpenUrlAbstract/FREE Full Text
  7. ↵
    Oddo, M., T. Renno, A. Attinger, T. Bakker, H. R. MacDonald, P. R. Meylan. 1998. Fas ligand-induced apoptosis of infected human macrophages reduces the viability of intracellular Mycobacterium tuberculosis. J. Immunol. 160: 5448
    OpenUrlAbstract/FREE Full Text
  8. ↵
    Wang, J., J. Wakeham, R. Harkness, Z. Xing. 1999. Macrophages are a significant source of type 1 cytokines during mycobacterial infection. J. Clin. Invest. 103: 1023
    OpenUrlCrossRefPubMed
  9. ↵
    Rook, G. A., R. Hernandez-Pando. 1996. The pathogenesis of tuberculosis. Annu. Rev. Microbiol. 50: 259
    OpenUrlCrossRefPubMed
  10. ↵
    Fenton, M. J., M. W. Vermeulen, S. Kim, M. Burdick, R. M. Strieter, H. Kornfeld. 1997. Induction of γ interferon production in human alveolar macrophages by Mycobacterium tuberculosis. Infect. Immun. 65: 5149
    OpenUrlAbstract/FREE Full Text
  11. ↵
    Flynn, J. L., J. Chan, K. J. Triebold, D. K. Dalton, T. A. Stewart, B. R. Bloom. 1993. An essential role for interferon γ in resistance to Mycobacterium tuberculosis infection. J. Exp. Med. 178: 2249
    OpenUrlAbstract/FREE Full Text
  12. ↵
    Cooper, A. M., D. K. Dalton, T. A. Stewart, J. P. Griffin, D. G. Russell, I. M. Orme. 1993. Disseminated tuberculosis in interferon γ gene-disrupted mice. J. Exp. Med. 178: 2243
    OpenUrlAbstract/FREE Full Text
  13. ↵
    Chan, J., S. H. E. Kaufmann. 1994. Immune mechanisms of protection. B. R. Bloom, ed. Tuberculosis: Pathogenesis, Protection, and Control 389-415. ASM Press, Washington, D.C.
  14. ↵
    Thepen, T., N. Van Rooijen, G. Kraal. 1989. Alveolar macrophage elimination in vivo is associated with an increase in pulmonary immune response in mice. J. Exp. Med. 170: 499
    OpenUrlAbstract/FREE Full Text
  15. ↵
    Naito, M., H. Nagai, S. Kawano, H. Umezu, H. Zhu, H. Moriyama, T. Yamamoto, H. Takatsuka, Y. Takei. 1996. Liposome-encapsulated dichloromethylene diphosphonate induces macrophage apoptosis in vivo and in vitro. J. Leukocyte Biol. 60: 337
    OpenUrlAbstract
  16. ↵
    Van Rooijen, N., A. Sanders, T. K. van den Berg. 1996. Apoptosis of macrophages induced by liposome-mediated intracellular delivery of clodronate and propamidine. J. Immunol. Methods 193: 93
    OpenUrlCrossRefPubMed
  17. ↵
    Van Rooijen, N., A. Sanders. 1994. Liposome mediated depletion of macrophages: mechanism of action, preparation of liposomes and applications. J. Immunol. Methods 174: 83
    OpenUrlCrossRefPubMed
  18. ↵
    Saunders, B. M., C. Cheers. 1996. Intranasal infection of beige mice with Mycobacterium avium complex: role of neutrophils and natural killer cells. Infect. Immun. 64: 4236
    OpenUrlAbstract/FREE Full Text
  19. ↵
    Erb, K. J., J. W. Holloway, A. Sobeck, H. Moll, G. Le Gros. 1998. Infection of mice with Mycobacterium bovis-bacillus Calmette-Guérin (BCG) suppresses allergen-induced airway eosinophilia. J. Exp. Med. 187: 561
    OpenUrlAbstract/FREE Full Text
  20. ↵
    Juffermans, N. P., S. Florquin, L. Camoglio, A. Verbon, A. H. Kolk, P. Speelman, S. J. van Deventer, T. van Der Poll. 2000. Interleukin-1 signaling is essential for host defense during murine pulmonary tuberculosis. J. Infect. Dis. 182: 902
    OpenUrlAbstract/FREE Full Text
  21. ↵
    Duriez, P. J., G. M. Shah. 1997. Cleavage of poly(ADP-ribose) polymerase: a sensitive parameter to study cell death. Biochem. Cell Biol. 75: 337
    OpenUrlCrossRefPubMed
  22. ↵
    Jordan, J. P., C. M. Hand, R. S. Markowitz, P. Black. 1992. Test for chemotherapeutic sensitivity of cerebral gliomas: use of colorimetric MTT assay. J. Neurooncol. 14: 19
    OpenUrlPubMed
  23. ↵
    Berg, J. T., S. T. Lee, T. Thepen, C. Y. Lee, M. F. Tsan. 1993. Depletion of alveolar macrophages by liposome-encapsulated dichloromethylene diphosphonate. J. Appl. Physiol. 74: 2812
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Kooguchi, K., S. Hashimoto, A. Kobayashi, Y. Kitamura, I. Kudoh, J. Wiener-Kronish, T. Sawa. 1998. Role of alveolar macrophages in initiation and regulation of inflammation in Pseudomonas aeruginosa pneumonia. Infect. Immun. 66: 3164
    OpenUrlAbstract/FREE Full Text
  25. ↵
    Cooper, A. M., J. L. Flynn. 1995. The protective immune response to Mycobacterium tuberculosis. Curr. Opin. Immunol. 7: 512
    OpenUrlCrossRefPubMed
  26. ↵
    Andersen, P.. 1997. Host responses and antigens involved in protective immunity to Mycobacterium tuberculosis. Scand. J. Immunol. 45: 115
    OpenUrlCrossRefPubMed
  27. ↵
    Caruso, A. M., N. Serbina, E. Klein, K. Triebold, B. R. Bloom, J. L. Flynn. 1999. Mice deficient in CD4 T cells have only transiently diminished levels of IFN-γ, yet succumb to tuberculosis. J. Immunol. 162: 5407
    OpenUrlAbstract/FREE Full Text
  28. ↵
    Antoni, B. A., P. Sabbatini, A. B. Rabson, E. White. 1995. Inhibition of apoptosis in human immunodeficiency virus-infected cells enhances virus production and facilitates persistent infection. J. Virol. 69: 2384
    OpenUrlAbstract/FREE Full Text
  29. ↵
    Vaux, D. L., G. Haecker, A. Strasser. 1994. An evolutionary perspective on apoptosis. Cell 76: 777
    OpenUrlCrossRefPubMed
  30. ↵
    Clem, R. J., L. K. Miller. 1993. Apoptosis reduces both the in vitro replication and the in vivo infectivity of a baculovirus. J. Virol. 67: 3730
    OpenUrlAbstract/FREE Full Text
  31. ↵
    Broug-Holub, E., G. B. Toews, J. F. van Iwaarden, R. M. Strieter, S. L. Kunkel, R. Paine, III, T. J. Standiford. 1997. Alveolar macrophages are required for protective pulmonary defenses in murine Klebsiella pneumonia: elimination of alveolar macrophages increases neutrophil recruitment but decreases bacterial clearance and survival. Infect. Immun. 65: 1139
    OpenUrlAbstract/FREE Full Text
  32. ↵
    Ottenhoff, T. H., T. Mutis. 1995. Role of cytotoxic cells in the protective immunity against and immunopathology of intracellular infections. Eur. J. Clin. Invest. 25: 371
    OpenUrlCrossRefPubMed
  33. ↵
    Cua, D. J., S. A. Stohlman. 1997. In vivo effects of T helper cell type 2 cytokines on macrophage antigen-presenting cell induction of T helper subsets. J. Immunol. 159: 5834
    OpenUrlAbstract
  34. ↵
    Goerdt, S., C. E. Orfanos. 1999. Other functions, other genes: alternative activation of antigen-presenting cells. Immunity 10: 137
    OpenUrlCrossRefPubMed
  35. ↵
    Martinez, J. A., T. E. King, Jr, K. Brown, C. A. Jennings, L. Borish, R. L. Mortenson, T. Z. Khan, T. W. Bost, D. W. Riches. 1997. Increased expression of the interleukin-10 gene by alveolar macrophages in interstitial lung disease. Am. J. Physiol. 273: L676
    OpenUrlAbstract/FREE Full Text
  36. ↵
    Magnan, A., D. van Pee, P. Bongrand, D. Vervloet. 1998. Alveolar macrophage interleukin (IL)-10 and IL-12 production in atopic asthma. Allergy 53: 1092
    OpenUrlCrossRefPubMed
  37. ↵
    Mosmann, T. R.. 1994. Properties and functions of interleukin-10. Adv. Immunol. 56: 1
    OpenUrlCrossRefPubMed
  38. ↵
    Stenger, S., R. L. Modlin. 1999. T cell mediated immunity to Mycobacterium tuberculosis. Curr. Opin. Microbiol. 2: 89
    OpenUrlCrossRefPubMed
  39. ↵
    Pedrosa, J., B. M. Saunders, R. Appelberg, I. M. Orme, M. T. Silva, A. M. Cooper. 2000. Neutrophils play a protective nonphagocytic role in systemic Mycobacterium tuberculosis infection of mice. Infect. Immun. 68: 577
    OpenUrlAbstract/FREE Full Text
  40. ↵
    Toews, G. B., W. C. Vial, M. M. Dunn, P. Guzzetta, G. Nunez, P. Stastny, M. F. Lipscomb. 1984. The accessory cell function of human alveolar macrophages in specific T cell proliferation. J. Immunol. 132: 181
    OpenUrlAbstract
  41. ↵
    Lipscomb, M. F., C. R. Lyons, G. Nunez, E. J. Ball, P. Stastny, W. Vial, V. Lem, J. Weissler, L. M. Miller. 1986. Human alveolar macrophages: HLA-DR-positive macrophages that are poor stimulators of a primary mixed leukocyte reaction. J. Immunol. 136: 497
    OpenUrlAbstract
  42. ↵
    Lyons, C. R., E. J. Ball, G. B. Toews, J. C. Weissler, P. Stastny, M. F. Lipscomb. 1986. Inability of human alveolar macrophages to stimulate resting T cells correlates with decreased antigen-specific T cell-macrophage binding. J. Immunol. 137: 1173
    OpenUrlAbstract
  43. ↵
    Kradin, R. L., K. M. McCarthy, C. I. Dailey, A. Burdeshaw, J. T. Kurnick, E. E. Schneeberger. 1987. The poor accessory cell function of macrophages in the rat may reflect their inability to form clusters with T cells. Clin. Immunol. Immunopathol. 44: 348
    OpenUrlCrossRefPubMed
  44. ↵
    Sertl, K., T. Takemura, E. Tschachler, V. J. Ferrans, M. A. Kaliner, E. M. Shevach. 1986. Dendritic cells with antigen-presenting capability reside in airway epithelium, lung parenchyma, and visceral pleura. J. Exp. Med. 163: 436
    OpenUrlAbstract/FREE Full Text
  45. ↵
    Masten, B. J., M. F. Lipscomb. 1999. Comparison of lung dendritic cells and B cells in stimulating naive antigen-specific T cells. J. Immunol. 162: 1310
    OpenUrlAbstract/FREE Full Text
  46. ↵
    Franke-Ullmann, G., C. Pfortner, P. Walter, C. Steinmuller, M. L. Lohmann-Matthes, L. Kobzik. 1996. Characterization of murine lung interstitial macrophages in comparison with alveolar macrophages in vitro. J. Immunol. 157: 3097
    OpenUrlAbstract
  47. ↵
    Holt, P. G.. 1979. Alveolar macrophages. II. Inhibition of lymphocyte proliferation by purified macrophages from rat lung. Immunology 37: 429
    OpenUrlPubMed
  48. ↵
    McCombs, C. C., J. P. Michalski, B. T. Westerfield, R. W. Light. 1982. Human alveolar macrophages suppress the proliferative response of peripheral blood lymphocytes. Chest 82: 266
    OpenUrlCrossRefPubMed
  49. ↵
    Bilyk, N., P. G. Holt. 1995. Cytokine modulation of the immunosuppressive phenotype of pulmonary alveolar macrophage populations. Immunology 86: 231
    OpenUrlPubMed
  50. ↵
    De Haan, A., G. Groen, J. Prop, N. van Rooijen, J. Wilschut. 1996. Mucosal immunoadjuvant activity of liposomes: role of alveolar macrophages. Immunology 89: 488
    OpenUrlCrossRefPubMed
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