Mycobacterium tuberculosis Growth Control by Lung Macrophages and CD8 Cells from Patient Contacts
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《美国呼吸和危急护理医学》
Department of Microbiology, Instituto Nacional de Enfermedades Respiratorias, Mexico City, Mexico
Department of Medicine and Center for Emerging Pathogens, University of Medicine and Dentistry of New Jersey, Newark, New Jersey
ABSTRACT
Rationale: Healthy household contacts (HHCs) of patients with active pulmonary tuberculosis are exposed aerogenically to Mycobacterium tuberculosis (Mtb), thus permitting the study of protective local immunity.
Objectives: To assess alveolar macrophage (AM) and autologous blood CD4 and CD8 T-cell–mediated Mtb growth control in HHCs and healthy, unexposed community control subjects (CCs).
Methods: AMs were infected with Mtb strains H37Ra and H37Rv at multiplicities of infection 0.1 and 1. Mtb colony-forming units were evaluated on Days 1, 4, and 7.
Main Results: CD8 T cells from HHCs in 1:1 cocultures with AMs significantly (p < 0.05) increased Mtb growth control by AMs. In CCs, no detectable contribution of CD8 T cells to Mtb growth control was observed. CD4 T cells did not increase Mtb growth control in HHCs or in CCs. IFN-, nitric oxide, and tumor necrosis factor were determined as potential mediators of Mtb growth control in AMs and AM/CD8 and AM/CD4 cocultures. IFN- production in AM/CD4 was twofold higher than that in AM/CD8 cocultures in both HHCs and CCs (p < 0.05). Nitric oxide production from AMs of HHCs increased on Days 4 and 7 and was undetectable in AMs from CCs. IFN- and nitric acid concentrations and Mtb growth control were not correlated. Tumor necrosis factor levels were significantly increased in AM/CD8 cocultures from HHCs compared with AM/CD8 cocultures from CCs (p < 0.05).
Conclusion: Aerogenic exposure to Mtb in HHCs leads to expansion of Mtb–specific effector CD8 T cells that limit Mtb growth in autologous AMs.
Key Words: interferon type II Mycobacterium tuberculosis nitric oxide T lymphocytes, effector macrophages, alveolar
The human lung is the primary site of respiratory uptake of aerosolized droplet nuclei containing Mycobacterium tuberculosis (Mtb) and of tuberculosis (TB) in the nonimmunocompromised host (1–4). Hence, the study of immune responses in the lung compartment provides a window into the fundamental biological interactions at the interface of Mtb with the host immune cells.
Epidemiologic studies suggest presence of human protective immunity against Mtb. The proportions of individuals that undergo primary infection with Mtb or reactivation disease after aerogenic Mtb exposure are usually low (3, 5, 6). Healthy household contacts (HHCs) of patients constitute a protected group, as only 5 to 10% of individuals exposed in households develop reactivation TB during their lifetime (3) and 50 to 75% do not acquire Mtb infection as determined by tuberculin skin testing (TST) (7, 8). By inference, it can thus be assumed that the local alveolar environment of HHC during its initial interaction with Mtb efficiently contains or abrogates the infection in the majority of cases despite sustained aerogenic Mtb exposure.
After the respiratory uptake of Mtb in aerosolized droplet nuclei, alveolar macrophages (AMs) are believed to be among the first cells to phagocytose, contain, and kill Mtb and to become targets of cytotoxic T-lymphocyte and CD8 effector T-cell activity (9–12). As mediators of innate and adaptive immune responses, Mtb-infected AMs release cytokines and chemokines (13–19). AMs probably are also stimulated in situ by cytokines such as IFN- and tumor necrosis factor (TNF-), which affect the bactericidal activity and control of Mtb (20).
There is general consensus that CD4 T cells play a pivotal role in human protective immunity against Mtb (21–23). CD4 T cells are the predominant source of IFN- and interleukin 2 (IL-2), capable of cytotoxicity against Mtb-infected target cells (24), and critical for the induction of protective-memory immunity, delayed-type hypersensitivity responses, and the development and maintenance of CD8 T-cell responses during Mtb infection (25).
CD8 T cells play an essential role as killer cells of Mtb-infected targets (26, 27). CD8 T cells lyse Mtb-infected macrophages via a Fas-independent granule exocytosis pathway (28–30) and the Fas–FasL interaction (12), which results in the apoptotic death of Mtb-infected target cells. Human peripheral CD8 T cells show Mtb specificity and recognize Mtb antigens, such as ESAT-6 (31), 19kD (32), and Mtb39 (33), in an HLA-1 dependent fashion (11, 31, 34–37) and contribute to the production of IFN- (38, 39) and TNF- on stimulation with Mtb (39).
IFN- and nitric oxide (NO) are key mediators of cell-mediated antimycobacterial activities. IFN- is crucial in the resistance to mycobacteria (40–43); its activity is mediated through the regulation of an array of genes that include reactive nitrogen intermediates. NO is released from Mtb-infected human blood monocytes (MNs) (44, 45) and AMs (46–48), confers antimycobacterial activity against Mtb (45, 48), and is present in human TB lung granulomas (49).
Our earlier studies in HHCs revealed increased frequencies of Mtb antigen 85–specific immune responses in peripheral blood mononuclear cells (PBMCs) (50) and bronchoalveolar cells (BACs) (51). These findings in BACs suggested induction of systemic and local protective-memory immunity in concurrently Mtb-exposed HHCs.
The current study assessed the Mtb growth–controlling capacity of AMs and of autologous blood CD4 and CD8 T cells from HHCs. Autologous CD8 T cells from HHCs significantly increased the capacity of in vitro–infected AMs to control the growth of Mtb. AM, AM/CD4, and AM/CD8 cocultures produced IFN- and NO after infection with Mtb; there was, however, no correlation between IFN- and NO concentrations and the growth control of Mtb conferred by AM and CD8 T cells. Some of the results of these studies have been previously reported in the form of abstracts (52–54).
METHODS
Study Subjects
HHCs of patients with active TB and CCs were screened for participation at the National Institute for Respiratory Diseases in Mexico City. Individuals willing to undergo bronchoalveolar lavage (BAL), venipuncture, HIV-1 serology, and TST were included in the study. A group of 13 HIV-1 seronegative HHCs (5 male, 8 female) were selected from first-grade relatives of acid-fast bacilli–positive patients with drug-sensitive TB. Each HHC shared the same house or room with an index patient for at least 3 mo before the patient's TB diagnosis. A group of 11 HIV-1 seronegative CCs without any known prior contact with patients with TB who had not been involved in patient care were recruited for the study. HHCs and CCs were 18 to 60 yr old. None of them had clinical or radiographic evidence of active pulmonary TB or of any other respiratory disease, or of acute, chronic, or immunocompromising diseases or therapies. Of the HHCs, eight were TST positive (> 10 mm), as were three CCs; the remaining HHCs and CCs were TST negative (< 10 mm).
Approval to obtain BACs by BAL and to perform venipunctures was given by the institutional review boards at the National Institute for Respiratory Diseases in Mexico City and the University of Medicine and Dentistry of New Jersey. Written, informed consent was obtained according to the guidelines of the U.S. Department of Health and Human Services.
Preparation of BACs
BACs were obtained by BAL as described previously (51). Briefly, after local anesthesia of the upper airways with 2% lidocaine solution and additional instillation of 1% lidocaine to prevent coughing, a flexible fiberoptic bronchoscope (P30; Olympus BF, New Hyde Park, NY) was wedged into the right middle lobe or the lingula. A total of 150 ml sterile 0.9% saline fluid was instilled in 20-ml aliquots into each of two subsegments. Mean yield of BAL fluid (BALF) was 78%. BALF was centrifuged at 400 x g for 15 min at 4°C. Pelleted BACs were resuspended in Roswell Park Memorial Institute (RPMI) 1640 (BioWhittaker, Walkersville, MD) with 50 μg/ml gentamicin sulfate, 200 mM L-glutamine, and 10% heat-inactivated pooled human serum (complete medium).
AMs were enriched from BACs using sheep red blood cells (SRBCs) (55) that bind to pan T-cell marker CD2. Briefly, 500 μl of washed SRBCs were activated with 1 IU neuraminidase (Sigma Chemical Co., Saint Louis, MO), and resuspended in RPMI. Then, 2 ml activated packed SRBCs and 2 ml fetal calf serum were added per milliliter of BACs (with 1–2 x 107 cells), and cell mixtures were incubated for 10 min at 37°C. Cells were then sedimented by centrifugation, incubated for 1 h on ice, and subsequently resuspended and overlaid on 3-ml Ficoll-Paque. After centrifugation, AMs were isolated from the buffy coat and washed three times. These cells were relatively lymphocyte-depleted and used as the AM population. Alveolar lymphocytes (ALs) were recovered from the pellet after lysis of the SRBCs with a buffer (0.01 M sodium bicarbonate–0.1 M ammonium chloride–0.1 mM ethylenediaminetetraacetic acid). The viability of BACs, AMs, and ALs was determined by exclusion of Trypan blue uptake, and was greater than 95% in both groups of subjects.
Preparation of PBMCs
Autologous PBMCs were prepared by density-gradient centrifugation of whole heparinized venous blood diluted 1:1 with RPMI over Ficoll-Paque (56) and resuspended in complete medium. The viability of PBMC was greater than 98% in each experiment from both groups of subjects.
Isolation of CD4 and CD8 Blood T Cells
Blood CD4 and CD8 T-cell subsets were prepared from PBMCs by stepwise immunomagnetic bead separation according to the manufacturer's instructions (Miltenyi Biotec, Auburn, CA). The mean purity of positively selected CD4 and CD8 T-cell preparations was 92 and 98%, respectively, as determined by fluorescence-activated cell sorting (FACS) analysis.
Preparation of H37Ra and H37Rv for In Vitro Infection
Suspensions of Mtb laboratory strains H37Ra and H37Rv (ATCC, Manassas, VA) were prepared in Middlebrook 7H9 broth medium (Difco Laboratories, Detroit, MI) (57). After a 21-d incubation at 37°C, mycobacterial stock solution was harvested. To prepare a suspension with single bacteria and disrupt mycobacterial clump formation, bacterial pellets were then resuspended in RPMI with 6% glycerol and vortexed for 5 min in the presence of five sterile 3-ml glass beads. The resulting suspension of mycobacteria was centrifuged at 900 x g for 10 min to remove any remaining large clumps. Supernatants with disaggregated mycobacterial stock cultures were then divided into aliquots and stored at –70°C until use. Mean concentrations of the frozen H37Ra and H37Rv stock suspensions were determined by counting colony-forming units (cfu) on 7H10 agar plates in triplicate serial dilutions of declumped stock suspensions between Days 21 and 28. This declumping procedure was performed in each experiment to ensure use of single-bacterial-cell suspensions and to establish the input amounts of bacteria for the infections at the various multiplicities of infection (MOI).
In Vitro Infection and Intracellular Growth of H37Ra and H37Rv in AMs
AMs were plated in triplicate wells at a concentration of 105 cells/well in 96-well round-bottom microtiter plates (Corning, Inc., Corning, NY) and were adhered and rested in RPMI with 10% pooled human serum (PHS) overnight at 37°C in 5% CO2. Cell-culture supernatants were carefully removed, and AMs were infected with H37Ra and H37Rv at MOI of 0.1 and 1, respectively. After a 1-h infection at 37°C in 5% CO2, culture supernatants were removed, avoiding dislodging of the cells. Plates were then washed three times to remove any extracellular mycobacteria. Complete medium (200 μl) was then added to the wells for further culturing of the infected AMs. Cocultures of AMs (105 AMs/well) with autologous blood CD4 or CD8 T cells (AM/CD4 and AM/CD8 cocultures) were prepared by adding purified CD4 or CD8 T cells at ratios of 1:1 to both uninfected and infected AMs. Cell-culture supernatants were removed on Days 0, 4, and 7 for subsequent determination of IFN-, NO, and TNF- by ELISA. AMs and AM/CD8 and AM/CD4 cocultures were then lysed with 0.1% sodium dodecyl sulfate (Sigma Chemical Co., St. Louis, MO) followed by addition of 7H9 and 20% bovine serum albumin (Sigma) on Days 0, 4, and 7. Serial dilutions of the lysates from triplicate wells from AMs and AM cocultures were then plated in triplicate onto Middlebrook 7H10 agar (Difco) and cfu were determined after a 21-d incubation at 37°C and 5% CO2. In representative experiments, viability of cell cultures was assessed on Days 0, 4, and 7 by exclusion of Trypan blue uptake.
Average viabilities of AMs and AM cocultures in four representative experiments from both groups of subjects were as follows: Day 0, 95%; Day 4, 95%; and Day 7, 93%.
Phagocytosis of Mtb by AMs was assessed in additional wells by identification of acid-fast bacilli with Ziehl-Neelsen staining of AMs after 1 h of infection at 37°C in 5% CO2 with H37Ra and H37Rv at MOI of 0.1 and 1. H37Ra and H37Rv phagocytosis at MOI 1 was similar (20–25%) for HHCs and CCs. At MOI 0.1, phagocytosis in HHCs was 11 to 15%, and in CCs it was 4%. Assessment of phagocytosis by cfu count on Day 0, however, revealed larger cfu numbers from HHC AMs than from CC AMs. The proportion of infected AMs was similar for HHCs and CCs; however, more bacteria had been phagocytosed by AMs from HHCs than by AMs from CCs.
NO Production
Culture supernatants from AMs and AM/CD4 and AM/CD8 cocultures were collected from 96-well plates at 1 h and on Days 4 and 7, divided into aliquots, and stored at –20°C until analysis. NO was quantified with the Griess reaction (58). Briefly, 50 μl of the culture supernatants were plated in 96-well flat-bottom plates and mixed with 50 μl of Griess reagent (0.1% naphtyl-ethilenediamine dihydrochloride and 1% sulfanilamide in 1 N HCl). The optical density was determined at 550 nm with a plate microreader (Labsystems Multiskan MCC/340; Labsystems, Helsinki, Finland). NaNO2 was used to generate a standard curve. NO2 concentrations in samples were expressed in μM/5 x 105 cells after background values with RPMI were subtracted.
IFN- and TNF- ELISA
Culture supernatants from AMs and AM/CD4 and AM/CD8 cocultures from Days 0, 4, and 7 were assayed for IFN- and TNF-. ELISA assays were performed using a final concentration of 1 μg/ml of antihuman IFN- (Endogen, Woburn, MA) or antihuman TNF- capture antibody (Pharmingen, San Diego, CA). Ninety-six-well plates were then blocked with 100 μl Superblock (Pierce, Rockford, IL) at room temperature (RT) for IFN- and at 37°C for TNF-. Standard curves with recombinant human IFN- (Endogen; 0–1,000 pg/ml) or TNF- (Endogen; 0–2,000 pg/ml) were made, and 50 μl of standard dilution or cell-culture supernatant was added per well followed by incubation for 2 h at 37°C. After washing, biotinylated secondary anti–IFN- antibody (Endogen; 0.25 μg/ml) or anti–TNF- antibody (Pharmingen; 0.5 μg/ml) was added and plates were incubated at RT for 1 h for IFN- and 45 min for TNF-. Plates were washed and streptavidin-phosphatase conjugate (Jackson Immunoresearch, West Grove, PA) was added to each well. Plates were incubated 30 min at RT in the dark and developed with phosphatase substrate (Sigma) for 20 min at RT in the dark. The reaction was stopped with 5% ethylenediaminetetraacetic acid. Absorbance was read on a microplate reader (Labsystems Multiskan MCC/340; Labsystems) at 450 nm. ELISA results are presented as mean pg/ml from triplicate wells.
Analysis of Cell Surface Markers by FACS
The subpopulations of BACs and the purity of enriched AMs from BACs and of blood CD4 and CD8 T cells were assessed by FACS analysis using fluorescein isothiocyanate–labeled anti-CD3, CD4, CD8, CD56, TCR, and CD14; phycoerythrin (PE)-labeled anti-CD69; and peridin-chlorophyll protein (PerCP)-labeled anti-CD3 antibodies. Fluorescein isothiocyanate–, PE-, and PerCP-labeled mouse anti-IgG1 and anti-IgG2a isotype antibodies were used to define quadrant positions in dot plots and percentage markers in histogram analyses. All antibodies were obtained from Becton Dickinson (San Jose, CA). Cells were antibody labeled, fixed with 1% paraformaldehyde, and kept at 4°C until acquisition in a FACScalibur (Becton Dickinson) within 24 h. Data analysis was done with the CELL-Quest software (Becton Dickinson). Cells were gated in the lymphocyte region (using FSC vs. SSC dot plot) to analyze the AL populations within the BACs. To assess the purity of AMs and of CD4 and CD8 blood T cells, no specific region (no gate) was selected.
Immunocytochemistry
Cellular profiles of BACs and AMs from HHCs and CCs on cytocentrifuge preparations were characterized using Wright's stain for nuclear morphology and a peroxidase stain that detects immature macrophages and neutrophils.
Statistical Analysis
Analyses to identify significant differences in cfu counts, IFN-, or NO concentrations between groups were done using a two-tailed Wilcoxon signed rank test, a nonparametric equivalent to a paired t test. Group means and SEM are presented. Statistical significance was set at p < 0.05, with all analyses being done using SPPS/PC+ version 11.5 (SPSS, Inc., Chicago, IL).
RESULTS
Subpopulations of BACs and Purity of AMs by Cytospin and FACS
By cytospin preparation, BACs from HHCs were 92% AMs and BACs from CCs were 90% AMs, with 8% ALs and less than 1% neutrophils; 5.5% of AMs from HHCs and 2% of AMs from CCs were peroxidase positive (p = 0.041). By FACS, ALs from HHCs were 47% CD3+CD4+, 22% CD3+CD8+, 9% CD56+, and 3% CD3+ TCR+. ALs from CCs were 42% CD3+CD4+, 29% CD3+CD8+, 10% CD56+, and 5% TCR+.
Purity of AM populations after SRBC rosetting was 91% in HHCs and 92% in CCs by cytospin preparation. Blood T-cell populations from HHCs and CCs were 92.5 to 95% and 91 to 94% CD3+CD4+ and CD3+CD8+, respectively.
HHCs
Antimycobacterial activity of AMs and AM/CD8 cocultures.
To test if concurrent aerogenic Mtb exposure of HHC induces bactericidal activity mediated by T cells on AMs, autologous blood CD8 and CD4 T cells from HHCs were added at a 1:1 ratio to in vitro Mtb-infected autologous AMs. Determinations of cfu from infections with H37Ra and H37Rv (MOI, 0.1 and 1) were made on Days 0, 4, and 7 in AMs and AM/CD8 and AM/CD4 cocultures.
Infection of AMs with H37Ra or H37Rv (MOI, 0.1) resulted in a steady cfu increase starting between 104 and 105 from Day 0 to Day 7 (Figures 1A and 1C). Interestingly, addition of CD8 T cells to AMs significantly increased control of H37Ra growth by AMs alone after 7 d. This observation was independent of the MOI (Figures 1A and 1B). CD8-mediated control of H37Rv growth was significant at MOI 0.1 (Figure 1C) but not at MOI 1 (Figure 1D).
There were no significant differences in the capacity of CD8 T cells from TST-negative and TST-positive donors to control Mtb growth.
In summary, CD8 T cells appear to increase Mtb growth control, particularly at MOI 0.1 and in H37Ra infections. The virulence of the H37Rv strain may have had an impact on the growth inhibition conferred by AMs and CD8 T cells as H37Rv cfu levels from AMs and AM/CD8 cultures were more than twofold greater than those from H37Ra-infected cells.
IFN- and NO production by AMs and AM/CD8 cocultures.
The importance of IFN- as an inducer of antimycobacterial activity of macrophages has been well established in the murine model. The current work assessed the production of IFN- in supernatants of AMs and AM/CD8 cocultures that were infected with H37Ra and H37Rv. AMs produced IFN- after in vitro infection with either Mtb strain. IFN- concentrations in H37Ra and H37Rv (MOI, 0.1) infection increased similarly from Day 0 to Day 7 (Figures 2A and 2C). AM infections with H37Rv and H37Ra at MOI 1 (Figures 2D and 2B, respectively) resulted in up to 30-fold increases from Day 0 to Day 7 (p < 0.05). IFN- levels of AMs were comparable at both MOI with H37Ra (Figures 2A and 2B).
IFN- concentrations in culture supernatants were affected by coculture of AMs with CD8 T cells, by the virulence of the infecting Mtb strain, and by the MOI.
IFN- concentrations in supernatants of infected AM/CD8 cocultures were one- to fourfold higher than that in AMs alone (p < 0.05).
IFN- levels in H37Rv-infected AMs and AM/CD8 cocultures were between two- and threefold higher than those in H37Ra-infected AMs and AM/CD8 cocultures (Figures 2A–2D). Regarding MOI, IFN- production was three- and twofold (p < 0.05) higher, respectively, in H37Ra and H37Rv infections with MOI 1 compared with infections with MOI 0.1 on Days 4 and 7. Constitutive IFN- levels in uninfected culture supernatants of AMs and AM/CD8 cocultures were low (means < 20 pg/ml) on Days 0, 4, and 7.
An important role for NO in the killing of Mtb and the significance of IFN- in the induction of NO production have clearly been established in the murine model. To assess in HHCs whether a relation existed between growth control of Mtb and NO, NO levels were measured in culture supernatants from AMs and AM/CD8 cocultures. AMs infected with H37Ra and H37Rv at MOI 0.1 produced low levels of NO (< 200 μM/105 cells); however, NO production was twofold higher in H37Rv- compared with H37Ra-infected AM cultures. NO levels increased from Day 0 to Day 7 in both strains at MOI 1 relative to MOI 0.1. NO production in AM/CD8 cocultures was lower than that in AM cultures alone (data not shown).
TNF- production in AMs and AM/CD8 cocultures.
TNF- levels were assessed in supernatants from AMs alone and from AM/CD8 cocultures. Median TNF- levels were twofold higher in AM/CD8 compared with AM cultures, indicating that CD8 T cells contribute to the production of TNF-. Levels of TNF- in AM/CD8 cultures were significantly higher in HHCs than in CCs (p < 0.05, data not shown). This observation suggests a role for TNF-–producing CD8 effector cells in the growth control of Mtb in this model.
Antimycobacterial activity and IFN- production in AMs and AM/CD4 cocultures.
In a subgroup of HHCs (n = 4), growth control of H37Ra and H37Rv was studied in AM/CD4 cocultures (Figure 3). The intracellular growth of H37Rv and H37Ra at MOI 0.1 in AMs and AM/CD4 cocultures is shown. There was no significant difference between cfu in AMs and AM/CD4 cocultures (Figures 3A and 3B) with either H37Ra or H37Rv, indicating lack of direct CD4 T-cell effects on Mtb growth in this model.
IFN- levels in AM/CD4 cocultures were higher compared with AM cultures both on infection with H37Ra and with H37Rv, and increased from Days 4 to 7 (Day 4, H37Ra, p < 0.05; Figures 3C and 3D). IFN- levels were also higher in AM/CD4 cocultures than in AM/CD8 cocultures, regardless of the infecting strain at MOI of 0.1 (not significant). Thus, as demonstrated by the lack of Mtb growth control conferred by CD4 T cells, increased IFN- production in AM/CD4 cocultures did not translate into improved growth control in AM/CD4 cocultures compared with AMs.
Healthy CCs
Antimycobacterial activity of AMs and AM/CD8 cocultures.
In a different set of experiments, contribution of peripheral CD4 and CD8 T cells to the growth control of H37Ra and H37Rv in AMs from CCs was studied. Infection of AMs and AM/CD4 and AM/CD8 cocultures with H37Ra (Figure 4A) and with H37Rv (Figure 4B) at MOI of 0.1 showed minimal contribution of CD4 and CD8 T cells to the growth control conferred by AMs alone. There were no significant differences in cfu among AMs and AM/CD4 and AM/CD8 cocultures. Similarly, no CD4 or CD8 T-cell effect on Mtb growth control in AMs was seen with either of the strains at MOI 1 (data not shown).
Concentrations of IFN- in AMs and AM/CD4 and AM/CD8 cocultures.
The production of IFN- in supernatants of AMs and AM/CD4 and AM/CD8 cocultures from CCs was also studied. Infection with H37Ra or H37Rv at MOI 0.1 (Figures 5A–5B) and MOI 1 (data not shown) induced similar increases of IFN- production by AMs from Day 0 to Day 7. As in HHCs, IFN- levels from AM/CD4 cocultures were significantly higher (p < 0.05) than those of AMs and AM/CD8 cocultures on Day 7 for all conditions: H37Ra and H37Rv at MOI 0.1 and MOI 1.
IFN- levels were higher in MOI 1 than in MOI 0.1 infections, regardless of the virulence of the strain (data not shown). Levels of constitutive IFN- production in uninfected AMs were low (< 50 pg/ml).
DISCUSSION
Healthy persons who are exposed to aerosolized Mtb allow for the study of early protective immunity in humans. To our knowledge, this study is the first of its kind to evaluate in HHCs the in vitro growth control of Mtb by ex vivo lung and blood cells. Growth control of avirulent H37Ra and virulent H37Rv by AMs alone and by AMs in coculture with autologous blood CD8 or CD4 T cells was assessed at the biologically relevant, low MOI of 0.1 and 1, which reduce mycobacterial clumping during infection and maximize AM viability in vitro.
Mtb growth in AM cultures from HHCs displayed a slow, steady increase of cfu over time. Addition of CD8 T cells to AM cultures, however, decreased H37Ra and H37Rv growth significantly. The Mtb growth-inhibiting effect of CD8 T cells on infected AMs was significantly larger than that conferred on AMs by CD4 T cells. Growth control was minimal when CD4 T cells were added to AM cultures in HHCs.
The greater Mtb growth control from CD8 compared with CD4 T cells in the current study contrasts with findings from Silver and colleagues (59). These investigators showed that CD4 T cells, rather than CD8 T cells, are responsible for growth control of H37Rv in human MNs. This discrepancy may be attributable to differences in the model systems with use of AMs in our study and of MNs in theirs. Also, differences in intensity, time points, duration of subjects' exposure to Mtb (subjects in this study were concurrently exposed to Mtb), and the capacity of AMs and MNs to phagocytose Mtb (57) may explain the observed dissimilar results.
Because the importance of CD8 T cells in protection against Mtb infection has been clearly demonstrated in murine models of infection, the present study builds on observations (31–33) that Mtb-specific CD8 T cells are expanded in the Mtb-infected human and that these cells play an important role in Mtb growth control in AMs.
Besides their cytotoxic activity against infected target cells such as AMs (11) or MNs (9, 10), CD8 T cells contribute to protective immunity by secreting cytokines such as IFN- and TNF- (12, 36, 60). To assess the concomitants of growth control of Mtb in this model system, the concentrations of IFN-, NO, and TNF- were measured in the supernatants of AM cultures and of AM/CD8 and AM/CD4 cocultures. Concentrations of IFN- in AM/CD8 cocultures were higher than in AMs alone but lower than in AM/CD4 cocultures. This finding corroborates earlier reports that showed that Mtb antigen-specific human CD4 T cells are the primary source of IFN- (34) and that CD8 T cells contribute, although less than CD4 cells, to the overall production of IFN- (34, 38). IFN- production detected in cultures from AMs alone may in part have been from contamination with ALs, which was similar between HHCs and CCs (8%).
IFN- levels in cocultures of AMs with CD4 and CD8 T cells in HHCs and CCs appeared to be unrelated to the growth control of Mtb when correlated with cfu. This mirrors earlier findings in BACs from patients with TB in whom alveolitis of activated T cells (61) increased levels of locally produced IFN-, and frequencies of Mtb antigen-specific IFN-–producing cells at the site of disease (55, 62, 63) accompanied unhindered progressive immunopathology.
NO was detectable in AM supernatants from HHCs on Days 4 and 7, corresponding with earlier reports of NO production by AMs (46, 47), or in lung granulomas of patients with TB (49, 64) or in healthy individuals (48, 65). However, as with IFN-, there was no evidence for a positive correlation between NO concentrations and growth control of H37Ra or H37Rv in this study. NO concentrations in this study were slightly higher in H37Ra than in H37Rv culture supernatants but unaffected by the MOI of the infection. Thus, although IFN- is believed to be an inducer of NO, the present study did not show a direct positive correlation between IFN- and NO concentrations in culture supernatants.
Interestingly, TNF- levels in culture supernatants were higher in AM/CD8 compared with AM cultures alone and higher in AM/CD8 cocultures from HHCs than AM/CD8 cocultures from CCs. These data suggest that TNF- from CD8 effector T cells (66) may contribute to the observed limiting effect of CD8 T cells on Mtb growth.
The present study also showed that the capacity of ex vivo AMs and CD8 T cells to control Mtb growth was affected by the virulence of the infecting Mtb strain. Levels of cfu from H37Ra-infected AMs were lower than those of H37Rv-infected AMs on Days 4 and 7 when compared under the same experimental conditions. These findings coincide with earlier observations (67) of increased cfu levels in human MN-derived macrophages that were infected with H37Rv or clinical isolates when compared with those infected with H37Ra.
CD8 T cells play an important role in human antimycobacterial immunity. Despite Mtb-induced CD4 responses (IFN- production), only CD8 T cells enhance mycobactericidal capacity of AMs in a uniform and significant manner in HHCs. The main reason for the improved capacity of CD8 T cells from HHCs to control Mtb growth may be the concurrent acute aerogenic exposure to Mtb and the subsequent expansion or activation of CD8 effector T cells. This notion is supported by a recent report of differential CD8 T-cell effector functions during acute and chronic phases of the immune response in the murine model. CD8 T cells isolated during the initial phase of infection showed lytic activity that is decreased in the chronic stage (68).
Recent evidence that human CD8 T cells preferentially recognize heavily infected cells (69) may suggest that differences in cfu levels of AMs from HHCs and CCs might have impinged on the CD8 T-cell effects on Mtb growth. However, differences in the model systems need to be considered. Lewinsohn studied infected dendritic cells that were exposed to CD8 T-cell clones and assessed IFN- release. In contrast, the current study assessed effects of ex vivo CD8-mediated control of Mtb growth in infected AMs that were not correlated directly with IFN- production, as IFN- production was similar in CCs and HHCs. Of interest also is the fact that CD8 T-cell activity controlling Mtb growth was lower in CCs than HHCs, although levels of cfu in AMs from CCs infected with MOI 1 (data not shown) were similar to those of AMs from HHCs infected with MOI 0.1. In vivo expansion and activation of CD8 T cells on Mtb exposure of HHCs thus contributed significantly to the observed differences between the study groups. However, during active Mtb infection, multiple factors including the degree of infection of antigen-presenting cells may determine expansion or activation of CD8 effector T cells.
In a set of separate studies from non–Mtb-exposed CCs, AMs were less permissive to Mtb infection because cfu levels were lower than those of AMs from HHCs. Differences in the activation status of AMs that resulted from concurrent in vivo exposure of HHCs to Mtb may be responsible for this finding. As in AMs from HHCs, Mtb cfu in AMs from CCs slowly increased between Day 0 and Day 4 with a plateau in growth between Day 4 and Day 7. However, no significant effect of CD8 or CD4 T cells was seen on this Mtb growth in AMs from CCs. These differences between HHCs and CCs were not due to variations in the input amount of bacteria or their preparation.
The limitations of this study are inherent in research on human subjects, particularly that involving the lung compartment. It is difficult to recruit large numbers of subjects and to control for extent and time point of exposure to Mtb in the household setting, which may result in variation of host immune responses among individuals. For small study groups, it may be explained that a statistically significant difference between TST-positive or TST-negative HHCs could not be detected. It is also important to consider that TST positivity in the study groups may have been due to both chronic Mtb infection or bacillus Calmette-Guerin vaccination (11 of 13 HHCs and 7 of 11 CCs had a bacillus Calmette-Guerin vaccination history) before the concurrent Mtb exposure period in the TB households. Processes leading to skin-test conversion may not translate directly into protection or improved effector T-cell activity. As suggested, the finding of CD8-mediated Mtb growth control that was observed here may be explained primarily by the recent Mtb exposure in the households. This exposure may have led to an expansion of Mtb-specific CD8 effector T cells, although additional mechanisms may have been at play. Longitudinal studies of larger cohorts of TB contacts may be required to assess immune responses in the lung in parallel with skin-test conversion and more specific Mtb antigen-based IFN- production assays.
In summary, together with earlier findings of increased protective-memory immunity in PBMCs (50) and BACs (51) of HHCs, this study shows that immune responses in HHCs are distinguishable from those of CCs. Furthermore, the presented ex vivo findings in HHCs suggest several processes that may occur in the intact host. First, Mtb-specific blood CD8 effector T cells may become expanded systemically after an exposure to Mtb or during ongoing aerogenic exposure. Second, these CD8 cells are capable of contributing to the control of Mtb growth in autologous Mtb-infected AMs. Third, recruitment and compartmentalization of the Mtb-specific CD8 T cells from the blood compartment to the local sites of inflammation in the lungs may play an important role in controlling the initial infection.
This work provides insights into human immunity that may require reinforcement with novel antituberculous vaccination or immunotherapeutic strategies. Antimycobacterial vaccination approaches that induce CD8 T-cell responses (70, 71) may be beneficial and particularly relevant in conditions of CD4-dependent immunodeficiencies.
Acknowledgments
The authors thank K. Bobadilla, M.D., and Leticia Sánchez, R.N., for screening family members of patients with TB and for screening individuals from the community. The authors are grateful for advice regarding statistical analysis from David Hom at the University of Medicine and Dentistry of New Jersey and grateful to Carmen Sarabia for support and careful review of the article. The authors acknowledge the help of the volunteers in this study.
FOOTNOTES
Supported by NHLBI 5 R01HL51630-10 and Conacyt 3479M.
Originally Published in Press as DOI: 10.1164/rccm.200503-411OC on October 6, 2005
Conflict of Interest Statement: None of the authors have a financial relationship with a commercial entity that has an interest in the subject of this manuscript.
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Department of Medicine and Center for Emerging Pathogens, University of Medicine and Dentistry of New Jersey, Newark, New Jersey
ABSTRACT
Rationale: Healthy household contacts (HHCs) of patients with active pulmonary tuberculosis are exposed aerogenically to Mycobacterium tuberculosis (Mtb), thus permitting the study of protective local immunity.
Objectives: To assess alveolar macrophage (AM) and autologous blood CD4 and CD8 T-cell–mediated Mtb growth control in HHCs and healthy, unexposed community control subjects (CCs).
Methods: AMs were infected with Mtb strains H37Ra and H37Rv at multiplicities of infection 0.1 and 1. Mtb colony-forming units were evaluated on Days 1, 4, and 7.
Main Results: CD8 T cells from HHCs in 1:1 cocultures with AMs significantly (p < 0.05) increased Mtb growth control by AMs. In CCs, no detectable contribution of CD8 T cells to Mtb growth control was observed. CD4 T cells did not increase Mtb growth control in HHCs or in CCs. IFN-, nitric oxide, and tumor necrosis factor were determined as potential mediators of Mtb growth control in AMs and AM/CD8 and AM/CD4 cocultures. IFN- production in AM/CD4 was twofold higher than that in AM/CD8 cocultures in both HHCs and CCs (p < 0.05). Nitric oxide production from AMs of HHCs increased on Days 4 and 7 and was undetectable in AMs from CCs. IFN- and nitric acid concentrations and Mtb growth control were not correlated. Tumor necrosis factor levels were significantly increased in AM/CD8 cocultures from HHCs compared with AM/CD8 cocultures from CCs (p < 0.05).
Conclusion: Aerogenic exposure to Mtb in HHCs leads to expansion of Mtb–specific effector CD8 T cells that limit Mtb growth in autologous AMs.
Key Words: interferon type II Mycobacterium tuberculosis nitric oxide T lymphocytes, effector macrophages, alveolar
The human lung is the primary site of respiratory uptake of aerosolized droplet nuclei containing Mycobacterium tuberculosis (Mtb) and of tuberculosis (TB) in the nonimmunocompromised host (1–4). Hence, the study of immune responses in the lung compartment provides a window into the fundamental biological interactions at the interface of Mtb with the host immune cells.
Epidemiologic studies suggest presence of human protective immunity against Mtb. The proportions of individuals that undergo primary infection with Mtb or reactivation disease after aerogenic Mtb exposure are usually low (3, 5, 6). Healthy household contacts (HHCs) of patients constitute a protected group, as only 5 to 10% of individuals exposed in households develop reactivation TB during their lifetime (3) and 50 to 75% do not acquire Mtb infection as determined by tuberculin skin testing (TST) (7, 8). By inference, it can thus be assumed that the local alveolar environment of HHC during its initial interaction with Mtb efficiently contains or abrogates the infection in the majority of cases despite sustained aerogenic Mtb exposure.
After the respiratory uptake of Mtb in aerosolized droplet nuclei, alveolar macrophages (AMs) are believed to be among the first cells to phagocytose, contain, and kill Mtb and to become targets of cytotoxic T-lymphocyte and CD8 effector T-cell activity (9–12). As mediators of innate and adaptive immune responses, Mtb-infected AMs release cytokines and chemokines (13–19). AMs probably are also stimulated in situ by cytokines such as IFN- and tumor necrosis factor (TNF-), which affect the bactericidal activity and control of Mtb (20).
There is general consensus that CD4 T cells play a pivotal role in human protective immunity against Mtb (21–23). CD4 T cells are the predominant source of IFN- and interleukin 2 (IL-2), capable of cytotoxicity against Mtb-infected target cells (24), and critical for the induction of protective-memory immunity, delayed-type hypersensitivity responses, and the development and maintenance of CD8 T-cell responses during Mtb infection (25).
CD8 T cells play an essential role as killer cells of Mtb-infected targets (26, 27). CD8 T cells lyse Mtb-infected macrophages via a Fas-independent granule exocytosis pathway (28–30) and the Fas–FasL interaction (12), which results in the apoptotic death of Mtb-infected target cells. Human peripheral CD8 T cells show Mtb specificity and recognize Mtb antigens, such as ESAT-6 (31), 19kD (32), and Mtb39 (33), in an HLA-1 dependent fashion (11, 31, 34–37) and contribute to the production of IFN- (38, 39) and TNF- on stimulation with Mtb (39).
IFN- and nitric oxide (NO) are key mediators of cell-mediated antimycobacterial activities. IFN- is crucial in the resistance to mycobacteria (40–43); its activity is mediated through the regulation of an array of genes that include reactive nitrogen intermediates. NO is released from Mtb-infected human blood monocytes (MNs) (44, 45) and AMs (46–48), confers antimycobacterial activity against Mtb (45, 48), and is present in human TB lung granulomas (49).
Our earlier studies in HHCs revealed increased frequencies of Mtb antigen 85–specific immune responses in peripheral blood mononuclear cells (PBMCs) (50) and bronchoalveolar cells (BACs) (51). These findings in BACs suggested induction of systemic and local protective-memory immunity in concurrently Mtb-exposed HHCs.
The current study assessed the Mtb growth–controlling capacity of AMs and of autologous blood CD4 and CD8 T cells from HHCs. Autologous CD8 T cells from HHCs significantly increased the capacity of in vitro–infected AMs to control the growth of Mtb. AM, AM/CD4, and AM/CD8 cocultures produced IFN- and NO after infection with Mtb; there was, however, no correlation between IFN- and NO concentrations and the growth control of Mtb conferred by AM and CD8 T cells. Some of the results of these studies have been previously reported in the form of abstracts (52–54).
METHODS
Study Subjects
HHCs of patients with active TB and CCs were screened for participation at the National Institute for Respiratory Diseases in Mexico City. Individuals willing to undergo bronchoalveolar lavage (BAL), venipuncture, HIV-1 serology, and TST were included in the study. A group of 13 HIV-1 seronegative HHCs (5 male, 8 female) were selected from first-grade relatives of acid-fast bacilli–positive patients with drug-sensitive TB. Each HHC shared the same house or room with an index patient for at least 3 mo before the patient's TB diagnosis. A group of 11 HIV-1 seronegative CCs without any known prior contact with patients with TB who had not been involved in patient care were recruited for the study. HHCs and CCs were 18 to 60 yr old. None of them had clinical or radiographic evidence of active pulmonary TB or of any other respiratory disease, or of acute, chronic, or immunocompromising diseases or therapies. Of the HHCs, eight were TST positive (> 10 mm), as were three CCs; the remaining HHCs and CCs were TST negative (< 10 mm).
Approval to obtain BACs by BAL and to perform venipunctures was given by the institutional review boards at the National Institute for Respiratory Diseases in Mexico City and the University of Medicine and Dentistry of New Jersey. Written, informed consent was obtained according to the guidelines of the U.S. Department of Health and Human Services.
Preparation of BACs
BACs were obtained by BAL as described previously (51). Briefly, after local anesthesia of the upper airways with 2% lidocaine solution and additional instillation of 1% lidocaine to prevent coughing, a flexible fiberoptic bronchoscope (P30; Olympus BF, New Hyde Park, NY) was wedged into the right middle lobe or the lingula. A total of 150 ml sterile 0.9% saline fluid was instilled in 20-ml aliquots into each of two subsegments. Mean yield of BAL fluid (BALF) was 78%. BALF was centrifuged at 400 x g for 15 min at 4°C. Pelleted BACs were resuspended in Roswell Park Memorial Institute (RPMI) 1640 (BioWhittaker, Walkersville, MD) with 50 μg/ml gentamicin sulfate, 200 mM L-glutamine, and 10% heat-inactivated pooled human serum (complete medium).
AMs were enriched from BACs using sheep red blood cells (SRBCs) (55) that bind to pan T-cell marker CD2. Briefly, 500 μl of washed SRBCs were activated with 1 IU neuraminidase (Sigma Chemical Co., Saint Louis, MO), and resuspended in RPMI. Then, 2 ml activated packed SRBCs and 2 ml fetal calf serum were added per milliliter of BACs (with 1–2 x 107 cells), and cell mixtures were incubated for 10 min at 37°C. Cells were then sedimented by centrifugation, incubated for 1 h on ice, and subsequently resuspended and overlaid on 3-ml Ficoll-Paque. After centrifugation, AMs were isolated from the buffy coat and washed three times. These cells were relatively lymphocyte-depleted and used as the AM population. Alveolar lymphocytes (ALs) were recovered from the pellet after lysis of the SRBCs with a buffer (0.01 M sodium bicarbonate–0.1 M ammonium chloride–0.1 mM ethylenediaminetetraacetic acid). The viability of BACs, AMs, and ALs was determined by exclusion of Trypan blue uptake, and was greater than 95% in both groups of subjects.
Preparation of PBMCs
Autologous PBMCs were prepared by density-gradient centrifugation of whole heparinized venous blood diluted 1:1 with RPMI over Ficoll-Paque (56) and resuspended in complete medium. The viability of PBMC was greater than 98% in each experiment from both groups of subjects.
Isolation of CD4 and CD8 Blood T Cells
Blood CD4 and CD8 T-cell subsets were prepared from PBMCs by stepwise immunomagnetic bead separation according to the manufacturer's instructions (Miltenyi Biotec, Auburn, CA). The mean purity of positively selected CD4 and CD8 T-cell preparations was 92 and 98%, respectively, as determined by fluorescence-activated cell sorting (FACS) analysis.
Preparation of H37Ra and H37Rv for In Vitro Infection
Suspensions of Mtb laboratory strains H37Ra and H37Rv (ATCC, Manassas, VA) were prepared in Middlebrook 7H9 broth medium (Difco Laboratories, Detroit, MI) (57). After a 21-d incubation at 37°C, mycobacterial stock solution was harvested. To prepare a suspension with single bacteria and disrupt mycobacterial clump formation, bacterial pellets were then resuspended in RPMI with 6% glycerol and vortexed for 5 min in the presence of five sterile 3-ml glass beads. The resulting suspension of mycobacteria was centrifuged at 900 x g for 10 min to remove any remaining large clumps. Supernatants with disaggregated mycobacterial stock cultures were then divided into aliquots and stored at –70°C until use. Mean concentrations of the frozen H37Ra and H37Rv stock suspensions were determined by counting colony-forming units (cfu) on 7H10 agar plates in triplicate serial dilutions of declumped stock suspensions between Days 21 and 28. This declumping procedure was performed in each experiment to ensure use of single-bacterial-cell suspensions and to establish the input amounts of bacteria for the infections at the various multiplicities of infection (MOI).
In Vitro Infection and Intracellular Growth of H37Ra and H37Rv in AMs
AMs were plated in triplicate wells at a concentration of 105 cells/well in 96-well round-bottom microtiter plates (Corning, Inc., Corning, NY) and were adhered and rested in RPMI with 10% pooled human serum (PHS) overnight at 37°C in 5% CO2. Cell-culture supernatants were carefully removed, and AMs were infected with H37Ra and H37Rv at MOI of 0.1 and 1, respectively. After a 1-h infection at 37°C in 5% CO2, culture supernatants were removed, avoiding dislodging of the cells. Plates were then washed three times to remove any extracellular mycobacteria. Complete medium (200 μl) was then added to the wells for further culturing of the infected AMs. Cocultures of AMs (105 AMs/well) with autologous blood CD4 or CD8 T cells (AM/CD4 and AM/CD8 cocultures) were prepared by adding purified CD4 or CD8 T cells at ratios of 1:1 to both uninfected and infected AMs. Cell-culture supernatants were removed on Days 0, 4, and 7 for subsequent determination of IFN-, NO, and TNF- by ELISA. AMs and AM/CD8 and AM/CD4 cocultures were then lysed with 0.1% sodium dodecyl sulfate (Sigma Chemical Co., St. Louis, MO) followed by addition of 7H9 and 20% bovine serum albumin (Sigma) on Days 0, 4, and 7. Serial dilutions of the lysates from triplicate wells from AMs and AM cocultures were then plated in triplicate onto Middlebrook 7H10 agar (Difco) and cfu were determined after a 21-d incubation at 37°C and 5% CO2. In representative experiments, viability of cell cultures was assessed on Days 0, 4, and 7 by exclusion of Trypan blue uptake.
Average viabilities of AMs and AM cocultures in four representative experiments from both groups of subjects were as follows: Day 0, 95%; Day 4, 95%; and Day 7, 93%.
Phagocytosis of Mtb by AMs was assessed in additional wells by identification of acid-fast bacilli with Ziehl-Neelsen staining of AMs after 1 h of infection at 37°C in 5% CO2 with H37Ra and H37Rv at MOI of 0.1 and 1. H37Ra and H37Rv phagocytosis at MOI 1 was similar (20–25%) for HHCs and CCs. At MOI 0.1, phagocytosis in HHCs was 11 to 15%, and in CCs it was 4%. Assessment of phagocytosis by cfu count on Day 0, however, revealed larger cfu numbers from HHC AMs than from CC AMs. The proportion of infected AMs was similar for HHCs and CCs; however, more bacteria had been phagocytosed by AMs from HHCs than by AMs from CCs.
NO Production
Culture supernatants from AMs and AM/CD4 and AM/CD8 cocultures were collected from 96-well plates at 1 h and on Days 4 and 7, divided into aliquots, and stored at –20°C until analysis. NO was quantified with the Griess reaction (58). Briefly, 50 μl of the culture supernatants were plated in 96-well flat-bottom plates and mixed with 50 μl of Griess reagent (0.1% naphtyl-ethilenediamine dihydrochloride and 1% sulfanilamide in 1 N HCl). The optical density was determined at 550 nm with a plate microreader (Labsystems Multiskan MCC/340; Labsystems, Helsinki, Finland). NaNO2 was used to generate a standard curve. NO2 concentrations in samples were expressed in μM/5 x 105 cells after background values with RPMI were subtracted.
IFN- and TNF- ELISA
Culture supernatants from AMs and AM/CD4 and AM/CD8 cocultures from Days 0, 4, and 7 were assayed for IFN- and TNF-. ELISA assays were performed using a final concentration of 1 μg/ml of antihuman IFN- (Endogen, Woburn, MA) or antihuman TNF- capture antibody (Pharmingen, San Diego, CA). Ninety-six-well plates were then blocked with 100 μl Superblock (Pierce, Rockford, IL) at room temperature (RT) for IFN- and at 37°C for TNF-. Standard curves with recombinant human IFN- (Endogen; 0–1,000 pg/ml) or TNF- (Endogen; 0–2,000 pg/ml) were made, and 50 μl of standard dilution or cell-culture supernatant was added per well followed by incubation for 2 h at 37°C. After washing, biotinylated secondary anti–IFN- antibody (Endogen; 0.25 μg/ml) or anti–TNF- antibody (Pharmingen; 0.5 μg/ml) was added and plates were incubated at RT for 1 h for IFN- and 45 min for TNF-. Plates were washed and streptavidin-phosphatase conjugate (Jackson Immunoresearch, West Grove, PA) was added to each well. Plates were incubated 30 min at RT in the dark and developed with phosphatase substrate (Sigma) for 20 min at RT in the dark. The reaction was stopped with 5% ethylenediaminetetraacetic acid. Absorbance was read on a microplate reader (Labsystems Multiskan MCC/340; Labsystems) at 450 nm. ELISA results are presented as mean pg/ml from triplicate wells.
Analysis of Cell Surface Markers by FACS
The subpopulations of BACs and the purity of enriched AMs from BACs and of blood CD4 and CD8 T cells were assessed by FACS analysis using fluorescein isothiocyanate–labeled anti-CD3, CD4, CD8, CD56, TCR, and CD14; phycoerythrin (PE)-labeled anti-CD69; and peridin-chlorophyll protein (PerCP)-labeled anti-CD3 antibodies. Fluorescein isothiocyanate–, PE-, and PerCP-labeled mouse anti-IgG1 and anti-IgG2a isotype antibodies were used to define quadrant positions in dot plots and percentage markers in histogram analyses. All antibodies were obtained from Becton Dickinson (San Jose, CA). Cells were antibody labeled, fixed with 1% paraformaldehyde, and kept at 4°C until acquisition in a FACScalibur (Becton Dickinson) within 24 h. Data analysis was done with the CELL-Quest software (Becton Dickinson). Cells were gated in the lymphocyte region (using FSC vs. SSC dot plot) to analyze the AL populations within the BACs. To assess the purity of AMs and of CD4 and CD8 blood T cells, no specific region (no gate) was selected.
Immunocytochemistry
Cellular profiles of BACs and AMs from HHCs and CCs on cytocentrifuge preparations were characterized using Wright's stain for nuclear morphology and a peroxidase stain that detects immature macrophages and neutrophils.
Statistical Analysis
Analyses to identify significant differences in cfu counts, IFN-, or NO concentrations between groups were done using a two-tailed Wilcoxon signed rank test, a nonparametric equivalent to a paired t test. Group means and SEM are presented. Statistical significance was set at p < 0.05, with all analyses being done using SPPS/PC+ version 11.5 (SPSS, Inc., Chicago, IL).
RESULTS
Subpopulations of BACs and Purity of AMs by Cytospin and FACS
By cytospin preparation, BACs from HHCs were 92% AMs and BACs from CCs were 90% AMs, with 8% ALs and less than 1% neutrophils; 5.5% of AMs from HHCs and 2% of AMs from CCs were peroxidase positive (p = 0.041). By FACS, ALs from HHCs were 47% CD3+CD4+, 22% CD3+CD8+, 9% CD56+, and 3% CD3+ TCR+. ALs from CCs were 42% CD3+CD4+, 29% CD3+CD8+, 10% CD56+, and 5% TCR+.
Purity of AM populations after SRBC rosetting was 91% in HHCs and 92% in CCs by cytospin preparation. Blood T-cell populations from HHCs and CCs were 92.5 to 95% and 91 to 94% CD3+CD4+ and CD3+CD8+, respectively.
HHCs
Antimycobacterial activity of AMs and AM/CD8 cocultures.
To test if concurrent aerogenic Mtb exposure of HHC induces bactericidal activity mediated by T cells on AMs, autologous blood CD8 and CD4 T cells from HHCs were added at a 1:1 ratio to in vitro Mtb-infected autologous AMs. Determinations of cfu from infections with H37Ra and H37Rv (MOI, 0.1 and 1) were made on Days 0, 4, and 7 in AMs and AM/CD8 and AM/CD4 cocultures.
Infection of AMs with H37Ra or H37Rv (MOI, 0.1) resulted in a steady cfu increase starting between 104 and 105 from Day 0 to Day 7 (Figures 1A and 1C). Interestingly, addition of CD8 T cells to AMs significantly increased control of H37Ra growth by AMs alone after 7 d. This observation was independent of the MOI (Figures 1A and 1B). CD8-mediated control of H37Rv growth was significant at MOI 0.1 (Figure 1C) but not at MOI 1 (Figure 1D).
There were no significant differences in the capacity of CD8 T cells from TST-negative and TST-positive donors to control Mtb growth.
In summary, CD8 T cells appear to increase Mtb growth control, particularly at MOI 0.1 and in H37Ra infections. The virulence of the H37Rv strain may have had an impact on the growth inhibition conferred by AMs and CD8 T cells as H37Rv cfu levels from AMs and AM/CD8 cultures were more than twofold greater than those from H37Ra-infected cells.
IFN- and NO production by AMs and AM/CD8 cocultures.
The importance of IFN- as an inducer of antimycobacterial activity of macrophages has been well established in the murine model. The current work assessed the production of IFN- in supernatants of AMs and AM/CD8 cocultures that were infected with H37Ra and H37Rv. AMs produced IFN- after in vitro infection with either Mtb strain. IFN- concentrations in H37Ra and H37Rv (MOI, 0.1) infection increased similarly from Day 0 to Day 7 (Figures 2A and 2C). AM infections with H37Rv and H37Ra at MOI 1 (Figures 2D and 2B, respectively) resulted in up to 30-fold increases from Day 0 to Day 7 (p < 0.05). IFN- levels of AMs were comparable at both MOI with H37Ra (Figures 2A and 2B).
IFN- concentrations in culture supernatants were affected by coculture of AMs with CD8 T cells, by the virulence of the infecting Mtb strain, and by the MOI.
IFN- concentrations in supernatants of infected AM/CD8 cocultures were one- to fourfold higher than that in AMs alone (p < 0.05).
IFN- levels in H37Rv-infected AMs and AM/CD8 cocultures were between two- and threefold higher than those in H37Ra-infected AMs and AM/CD8 cocultures (Figures 2A–2D). Regarding MOI, IFN- production was three- and twofold (p < 0.05) higher, respectively, in H37Ra and H37Rv infections with MOI 1 compared with infections with MOI 0.1 on Days 4 and 7. Constitutive IFN- levels in uninfected culture supernatants of AMs and AM/CD8 cocultures were low (means < 20 pg/ml) on Days 0, 4, and 7.
An important role for NO in the killing of Mtb and the significance of IFN- in the induction of NO production have clearly been established in the murine model. To assess in HHCs whether a relation existed between growth control of Mtb and NO, NO levels were measured in culture supernatants from AMs and AM/CD8 cocultures. AMs infected with H37Ra and H37Rv at MOI 0.1 produced low levels of NO (< 200 μM/105 cells); however, NO production was twofold higher in H37Rv- compared with H37Ra-infected AM cultures. NO levels increased from Day 0 to Day 7 in both strains at MOI 1 relative to MOI 0.1. NO production in AM/CD8 cocultures was lower than that in AM cultures alone (data not shown).
TNF- production in AMs and AM/CD8 cocultures.
TNF- levels were assessed in supernatants from AMs alone and from AM/CD8 cocultures. Median TNF- levels were twofold higher in AM/CD8 compared with AM cultures, indicating that CD8 T cells contribute to the production of TNF-. Levels of TNF- in AM/CD8 cultures were significantly higher in HHCs than in CCs (p < 0.05, data not shown). This observation suggests a role for TNF-–producing CD8 effector cells in the growth control of Mtb in this model.
Antimycobacterial activity and IFN- production in AMs and AM/CD4 cocultures.
In a subgroup of HHCs (n = 4), growth control of H37Ra and H37Rv was studied in AM/CD4 cocultures (Figure 3). The intracellular growth of H37Rv and H37Ra at MOI 0.1 in AMs and AM/CD4 cocultures is shown. There was no significant difference between cfu in AMs and AM/CD4 cocultures (Figures 3A and 3B) with either H37Ra or H37Rv, indicating lack of direct CD4 T-cell effects on Mtb growth in this model.
IFN- levels in AM/CD4 cocultures were higher compared with AM cultures both on infection with H37Ra and with H37Rv, and increased from Days 4 to 7 (Day 4, H37Ra, p < 0.05; Figures 3C and 3D). IFN- levels were also higher in AM/CD4 cocultures than in AM/CD8 cocultures, regardless of the infecting strain at MOI of 0.1 (not significant). Thus, as demonstrated by the lack of Mtb growth control conferred by CD4 T cells, increased IFN- production in AM/CD4 cocultures did not translate into improved growth control in AM/CD4 cocultures compared with AMs.
Healthy CCs
Antimycobacterial activity of AMs and AM/CD8 cocultures.
In a different set of experiments, contribution of peripheral CD4 and CD8 T cells to the growth control of H37Ra and H37Rv in AMs from CCs was studied. Infection of AMs and AM/CD4 and AM/CD8 cocultures with H37Ra (Figure 4A) and with H37Rv (Figure 4B) at MOI of 0.1 showed minimal contribution of CD4 and CD8 T cells to the growth control conferred by AMs alone. There were no significant differences in cfu among AMs and AM/CD4 and AM/CD8 cocultures. Similarly, no CD4 or CD8 T-cell effect on Mtb growth control in AMs was seen with either of the strains at MOI 1 (data not shown).
Concentrations of IFN- in AMs and AM/CD4 and AM/CD8 cocultures.
The production of IFN- in supernatants of AMs and AM/CD4 and AM/CD8 cocultures from CCs was also studied. Infection with H37Ra or H37Rv at MOI 0.1 (Figures 5A–5B) and MOI 1 (data not shown) induced similar increases of IFN- production by AMs from Day 0 to Day 7. As in HHCs, IFN- levels from AM/CD4 cocultures were significantly higher (p < 0.05) than those of AMs and AM/CD8 cocultures on Day 7 for all conditions: H37Ra and H37Rv at MOI 0.1 and MOI 1.
IFN- levels were higher in MOI 1 than in MOI 0.1 infections, regardless of the virulence of the strain (data not shown). Levels of constitutive IFN- production in uninfected AMs were low (< 50 pg/ml).
DISCUSSION
Healthy persons who are exposed to aerosolized Mtb allow for the study of early protective immunity in humans. To our knowledge, this study is the first of its kind to evaluate in HHCs the in vitro growth control of Mtb by ex vivo lung and blood cells. Growth control of avirulent H37Ra and virulent H37Rv by AMs alone and by AMs in coculture with autologous blood CD8 or CD4 T cells was assessed at the biologically relevant, low MOI of 0.1 and 1, which reduce mycobacterial clumping during infection and maximize AM viability in vitro.
Mtb growth in AM cultures from HHCs displayed a slow, steady increase of cfu over time. Addition of CD8 T cells to AM cultures, however, decreased H37Ra and H37Rv growth significantly. The Mtb growth-inhibiting effect of CD8 T cells on infected AMs was significantly larger than that conferred on AMs by CD4 T cells. Growth control was minimal when CD4 T cells were added to AM cultures in HHCs.
The greater Mtb growth control from CD8 compared with CD4 T cells in the current study contrasts with findings from Silver and colleagues (59). These investigators showed that CD4 T cells, rather than CD8 T cells, are responsible for growth control of H37Rv in human MNs. This discrepancy may be attributable to differences in the model systems with use of AMs in our study and of MNs in theirs. Also, differences in intensity, time points, duration of subjects' exposure to Mtb (subjects in this study were concurrently exposed to Mtb), and the capacity of AMs and MNs to phagocytose Mtb (57) may explain the observed dissimilar results.
Because the importance of CD8 T cells in protection against Mtb infection has been clearly demonstrated in murine models of infection, the present study builds on observations (31–33) that Mtb-specific CD8 T cells are expanded in the Mtb-infected human and that these cells play an important role in Mtb growth control in AMs.
Besides their cytotoxic activity against infected target cells such as AMs (11) or MNs (9, 10), CD8 T cells contribute to protective immunity by secreting cytokines such as IFN- and TNF- (12, 36, 60). To assess the concomitants of growth control of Mtb in this model system, the concentrations of IFN-, NO, and TNF- were measured in the supernatants of AM cultures and of AM/CD8 and AM/CD4 cocultures. Concentrations of IFN- in AM/CD8 cocultures were higher than in AMs alone but lower than in AM/CD4 cocultures. This finding corroborates earlier reports that showed that Mtb antigen-specific human CD4 T cells are the primary source of IFN- (34) and that CD8 T cells contribute, although less than CD4 cells, to the overall production of IFN- (34, 38). IFN- production detected in cultures from AMs alone may in part have been from contamination with ALs, which was similar between HHCs and CCs (8%).
IFN- levels in cocultures of AMs with CD4 and CD8 T cells in HHCs and CCs appeared to be unrelated to the growth control of Mtb when correlated with cfu. This mirrors earlier findings in BACs from patients with TB in whom alveolitis of activated T cells (61) increased levels of locally produced IFN-, and frequencies of Mtb antigen-specific IFN-–producing cells at the site of disease (55, 62, 63) accompanied unhindered progressive immunopathology.
NO was detectable in AM supernatants from HHCs on Days 4 and 7, corresponding with earlier reports of NO production by AMs (46, 47), or in lung granulomas of patients with TB (49, 64) or in healthy individuals (48, 65). However, as with IFN-, there was no evidence for a positive correlation between NO concentrations and growth control of H37Ra or H37Rv in this study. NO concentrations in this study were slightly higher in H37Ra than in H37Rv culture supernatants but unaffected by the MOI of the infection. Thus, although IFN- is believed to be an inducer of NO, the present study did not show a direct positive correlation between IFN- and NO concentrations in culture supernatants.
Interestingly, TNF- levels in culture supernatants were higher in AM/CD8 compared with AM cultures alone and higher in AM/CD8 cocultures from HHCs than AM/CD8 cocultures from CCs. These data suggest that TNF- from CD8 effector T cells (66) may contribute to the observed limiting effect of CD8 T cells on Mtb growth.
The present study also showed that the capacity of ex vivo AMs and CD8 T cells to control Mtb growth was affected by the virulence of the infecting Mtb strain. Levels of cfu from H37Ra-infected AMs were lower than those of H37Rv-infected AMs on Days 4 and 7 when compared under the same experimental conditions. These findings coincide with earlier observations (67) of increased cfu levels in human MN-derived macrophages that were infected with H37Rv or clinical isolates when compared with those infected with H37Ra.
CD8 T cells play an important role in human antimycobacterial immunity. Despite Mtb-induced CD4 responses (IFN- production), only CD8 T cells enhance mycobactericidal capacity of AMs in a uniform and significant manner in HHCs. The main reason for the improved capacity of CD8 T cells from HHCs to control Mtb growth may be the concurrent acute aerogenic exposure to Mtb and the subsequent expansion or activation of CD8 effector T cells. This notion is supported by a recent report of differential CD8 T-cell effector functions during acute and chronic phases of the immune response in the murine model. CD8 T cells isolated during the initial phase of infection showed lytic activity that is decreased in the chronic stage (68).
Recent evidence that human CD8 T cells preferentially recognize heavily infected cells (69) may suggest that differences in cfu levels of AMs from HHCs and CCs might have impinged on the CD8 T-cell effects on Mtb growth. However, differences in the model systems need to be considered. Lewinsohn studied infected dendritic cells that were exposed to CD8 T-cell clones and assessed IFN- release. In contrast, the current study assessed effects of ex vivo CD8-mediated control of Mtb growth in infected AMs that were not correlated directly with IFN- production, as IFN- production was similar in CCs and HHCs. Of interest also is the fact that CD8 T-cell activity controlling Mtb growth was lower in CCs than HHCs, although levels of cfu in AMs from CCs infected with MOI 1 (data not shown) were similar to those of AMs from HHCs infected with MOI 0.1. In vivo expansion and activation of CD8 T cells on Mtb exposure of HHCs thus contributed significantly to the observed differences between the study groups. However, during active Mtb infection, multiple factors including the degree of infection of antigen-presenting cells may determine expansion or activation of CD8 effector T cells.
In a set of separate studies from non–Mtb-exposed CCs, AMs were less permissive to Mtb infection because cfu levels were lower than those of AMs from HHCs. Differences in the activation status of AMs that resulted from concurrent in vivo exposure of HHCs to Mtb may be responsible for this finding. As in AMs from HHCs, Mtb cfu in AMs from CCs slowly increased between Day 0 and Day 4 with a plateau in growth between Day 4 and Day 7. However, no significant effect of CD8 or CD4 T cells was seen on this Mtb growth in AMs from CCs. These differences between HHCs and CCs were not due to variations in the input amount of bacteria or their preparation.
The limitations of this study are inherent in research on human subjects, particularly that involving the lung compartment. It is difficult to recruit large numbers of subjects and to control for extent and time point of exposure to Mtb in the household setting, which may result in variation of host immune responses among individuals. For small study groups, it may be explained that a statistically significant difference between TST-positive or TST-negative HHCs could not be detected. It is also important to consider that TST positivity in the study groups may have been due to both chronic Mtb infection or bacillus Calmette-Guerin vaccination (11 of 13 HHCs and 7 of 11 CCs had a bacillus Calmette-Guerin vaccination history) before the concurrent Mtb exposure period in the TB households. Processes leading to skin-test conversion may not translate directly into protection or improved effector T-cell activity. As suggested, the finding of CD8-mediated Mtb growth control that was observed here may be explained primarily by the recent Mtb exposure in the households. This exposure may have led to an expansion of Mtb-specific CD8 effector T cells, although additional mechanisms may have been at play. Longitudinal studies of larger cohorts of TB contacts may be required to assess immune responses in the lung in parallel with skin-test conversion and more specific Mtb antigen-based IFN- production assays.
In summary, together with earlier findings of increased protective-memory immunity in PBMCs (50) and BACs (51) of HHCs, this study shows that immune responses in HHCs are distinguishable from those of CCs. Furthermore, the presented ex vivo findings in HHCs suggest several processes that may occur in the intact host. First, Mtb-specific blood CD8 effector T cells may become expanded systemically after an exposure to Mtb or during ongoing aerogenic exposure. Second, these CD8 cells are capable of contributing to the control of Mtb growth in autologous Mtb-infected AMs. Third, recruitment and compartmentalization of the Mtb-specific CD8 T cells from the blood compartment to the local sites of inflammation in the lungs may play an important role in controlling the initial infection.
This work provides insights into human immunity that may require reinforcement with novel antituberculous vaccination or immunotherapeutic strategies. Antimycobacterial vaccination approaches that induce CD8 T-cell responses (70, 71) may be beneficial and particularly relevant in conditions of CD4-dependent immunodeficiencies.
Acknowledgments
The authors thank K. Bobadilla, M.D., and Leticia Sánchez, R.N., for screening family members of patients with TB and for screening individuals from the community. The authors are grateful for advice regarding statistical analysis from David Hom at the University of Medicine and Dentistry of New Jersey and grateful to Carmen Sarabia for support and careful review of the article. The authors acknowledge the help of the volunteers in this study.
FOOTNOTES
Supported by NHLBI 5 R01HL51630-10 and Conacyt 3479M.
Originally Published in Press as DOI: 10.1164/rccm.200503-411OC on October 6, 2005
Conflict of Interest Statement: None of the authors have a financial relationship with a commercial entity that has an interest in the subject of this manuscript.
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