The control of Mycobacterium tuberculosis infection is dependent on the development of an adaptive immune response, which is mediated by granulomas. The granuloma is a dynamic structure that forms in the lung and consists primarily of macrophages and lymphocytes. For this structure to be effective in containment of the bacillus, it must develop in an organized and timely manner. The formation of the granuloma is dependent on recruitment of activated cells through adhesion molecules and chemokines. M. tuberculosis infection causes an increase in the expression of β-chemokines CCL3, CCL4, and CCL5, and their receptor CCR5, in the lungs. In this study, we demonstrate that CCR5-transgenic knockout mice were capable of recruiting immune cells to the lung to form granulomas. CCR5−/− mice successfully induced a Th1 response and controlled infection. Unexpectedly, M. tuberculosis infection in these mice resulted in greater numbers of lymphocytes migrating to the lung and higher levels of many inflammatory cytokines, compared with wild-type mice, without apparent long-term detrimental effects. In the absence of CCR5, there were more dendritic cells in the lung-draining lymph nodes and more primed T lymphocytes in these mice. Bacterial numbers in the lymph nodes were also higher in CCR5−/− mice. Therefore, CCR5 may play a role in the migration of dendritic cells to and from the lymph nodes during M. tuberculosis infection.

Recent epidemiological studies estimate that one-third of the world’s population is infected with Mycobacterium tuberculosis (1). A majority of individuals exposed to M. tuberculosis never exhibit clinical disease. Rather, they mount a protective immune response to the infection. The protective response, which is induced during initial infection, requires CD4+ lymphocytes, CD8+ lymphocytes, the Th1-type cytokines IFN-γ and TNF, and activated macrophages (reviewed in Ref. 2). The cooperation between the cells and cytokines requires close interaction, which is achieved following migration and granuloma formation in the lungs. The hallmark of infection in the lung is granuloma formation. The granuloma structure consists of clusters of macrophages, B and T lymphocytes, and dendritic cells (DCs)3 (3, 4, 5). It physically contains the mycobacteria and creates a microenvironment for immune cell interaction, limiting M. tuberculosis growth and dissemination.

When activated by APCs, M. tuberculosis-specific CD4+ and CD8+ lymphocytes produce IFN-γ and TNF (6, 7, 8). IFN-γ then acts along with a second signal, such as TNF or mycobacterial Ags, to activate macrophages and DCs. Macrophage activation can lead to killing of M. tuberculosis through the production of reactive nitrogen intermediates and acidification of the phagolysosome (9). DC activation does not kill M. tuberculosis, but limits growth of the organism (10, 11). This may allow the DCs to function more effectively as APCs in the lymph nodes. CD8+ lymphocytes may participate in the protective response against tuberculosis not only through production of cytokines but also by functioning as CTL (reviewed in Ref. 12). Determining the factors responsible for bringing the immune cells to the lungs and organizing the cells into a granuloma is important for understanding the progression and control of M. tuberculosis infection.

Cell migration to sites of infection is mediated not only by adhesion molecules but also by chemotactic cytokines, chemokines. Chemokines influence cell migration and activation through signaling initiated by binding to specific G protein-coupled receptors. There is redundancy in the chemokine system. Specific to this study, chemokines CCL3, CCL4, and CCL5 are ligands of CCR5 (13). However, CCL4 and CCL5 can also signal through CCR1 and CCR3. CCR1 is expressed on Th1 lymphocytes, neutrophils, monocytes, B cells, and immature DCs, whereas CCR3 is expressed on Th2 cells, eosinophils, and basophils. Chemokine and chemokine receptor expression has been studied to a limited degree in M. tuberculosis (reviewed in Ref. 14). In murine models, gene expression of α and β chemokines has been detected in the lungs following M. tuberculosis infection (15, 16, 17). In this study, we addressed the role of CCR5 in murine M. tuberculosis infection.

CCR5 is a coreceptor for HIV (18), and as a result, an interest in the role of CCR5 during the immune response to infectious diseases and as a target for HIV therapy has emerged. CCR5 is expressed on granulocytes, macrophages, immature DCs, and CD8+ lymphocytes, and at a high level on Th1 lymphocytes (13, 19). Studies in infectious disease models illustrate the variety of functions of CCR5 in the immune response. In CCR5−/− mice infected with Leishmania donovani, the boundaries of the granulomas were difficult to demarcate, yet the mice had fewer parasites than wild-type mice (20). In contrast, CCR5−/− mice were more susceptible to influenza A virus, Cryptococcus neoformans, Listeria monocytogenes, and Toxoplasma gondii. CCR5 was associated with elimination of extracellular polysaccharide during C. neoformans infection (21), induction of IL-12 during T. gondii infection (22), and trafficking of cells to sites of infection (23, 24). Understanding the role of CCR5 in each infection may enhance the development of antimicrobial drugs and vaccines.

CCR5−/− mice controlled M. tuberculosis growth in the lungs, spleen, and liver, but had higher bacterial loads in the lung-draining lymph nodes. Substantially higher numbers of T lymphocytes were found in the lungs of CCR5−/− mice, compared with controls. The data presented in this study support a potential role for CCR5 in DC migration. Our data are consistent with the hypotheses that, in the absence of CCR5, higher bacterial loads in the lymph nodes are due to more DCs carrying M. tuberculosis from the lungs to the lymph nodes or there is retention of DCs in the lymph nodes presenting Ag, resulting in more priming of lymphocytes in the lymph nodes. These primed/activated lymphocytes then migrate to the site of infection (the lungs). These results suggest that CCR5 may be important for the migration to and retention of DCs in lymph nodes during infection.

B6;129PF2/J (The Jackson Laboratory, Bar Harbor, ME) or C57BL/6 (The Jackson Laboratory) and CCR5−/− (B6;129P-Cmkbr5tm1Kuz; The Jackson Laboratory) male and female mice (8–14 wk old) were used in all experiments. CCR5−/− mice were bred at the University of Pittsburgh in specific pathogen-free facilities. There were no differences observed with infection of B6;129PF2/J mice compared with C57BL/6 mice. All experiments showing CFU, lung and lymph node cell numbers, and histological analysis were performed comparing CCR5−/− mice with both C57BL/6 and B6;129PF2/J mice. In this paper, we are presenting the data with only B6;129PF2 mice for those assays just described. The other assays performed were with C57BL/6 mice as the controls. All infected mice were maintained in the biosafety level 3 animal laboratories and routinely monitored for murine pathogens. The University Institutional Animal Care and Use Committee approved all animal protocols used in this study.

All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) unless otherwise noted. Middlebrook 7H9 liquid medium and 7H10 agar were obtained from Difco Laboratories (Detroit, MI). Abs used in flow cytometry were obtained from BD Pharmingen (San Diego, CA).

To prepare bacterial stock, M. tuberculosis strain Erdman (Trudeau Institute, Saranac Lake, NY) was used to infect mice, and then bacteria were harvested from their lungs, expanded in 7H9 liquid medium, and stored in aliquots at −80°C. Mice were infected via the aerosol route using a nose-only exposure unit (InTox Products, Albuquerque, NM) and exposed to M. tuberculosis (1 × 107 CFU/ml in the nebulizer chamber) for 20 min, followed by 5 min of air. This resulted in reproducible delivery of 50–100 viable CFU of M. tuberculosis as described previously (25), which was confirmed by CFU determination on the lungs of two to three infected mice 1 day postinfection. Mice were infected i.v. via tail vein with 1 × 104 CFU in 100 μl of PBS/0.05% Tween 80. The tissue bacillary load was quantified by plating serial dilutions of the lung, liver, and spleen homogenates onto 7H10 agar as described previously (26).

Macrophages were derived from the C57BL/6 mice bone marrow based on adherence. The mice were euthanized, and bone marrow was flushed out of the femur and tibia bones with DMEM as previously described (10). The bone marrow suspension was washed twice with 2% FBS in PBS, and the cells were counted. A total of 2 × 106 cells was plated on non-tissue culture-treated Lab Tek petri dishes (Sigma-Aldrich) in 25 ml of macrophage medium (25% L cell supernatant, 20% FBS, 1% l-glutamine, 1% pyruvate, and 1% nonessential amino acids). After 4 days in culture, the cells were fed with 10 ml of fresh macrophage medium. On day 6, macrophages were infected with M. tuberculosis (multiplicity of infection (MOI), 4). Four hours postinfection, the supernatants were removed, cells were washed, and fresh medium was added.

DCs are derived from the C57BL/6 mice bone marrow. The bone marrow was obtained as described as above. After the cells were washed and counted, the cells were resuspended in two non-tissue culture-treated deep dish petri dishes in 20 ml/plate DC medium (10% FBS, 1% sodium pyruvate, and 1% l-glutamate) overnight. The following day, the nonadherent and semiadherent cells were removed from the plates, spun down, and cultured at 1 × 106 cells/ml in DC medium with 20 ng/ml GM-CSF and 20 ng/ml IL-4 (PeproTech, Rocky Hill, NJ) in p75 filter flasks. On day 3 of culture, the cells were fed with an additional 5 ml of DC medium and 100 ng of GM-CSF and IL-4. DCs were infected with M. tuberculosis on day 6 of culture (MOI, 4) as previously described (10).

To determine cellular infiltrate in the lung, at 10-day intervals lungs were removed for flow cytometric analysis. A single-cell suspension of the lungs or mediastinal lymph node was prepared by pushing the tissue through a cell strainer (Fisher, Pittsburgh, PA) as previously described (27). RBC were lysed with lysis buffer (0.144 M NH4Cl/0.017 M Tris (pH = 7.65)) and washed, and then the single-cell suspension was counted. The samples were stained with 0.2 μg of anti-CD4, anti-CD8, and anti-CD69, or 0.15 μg of anti-Gr1, 0.2 μg of anti-CD11c, and 0.2 μg of anti-CD11b in FACS buffer (0.1% Na azide, 0.1% BSA, and 20% mouse serum). Following washes, the cells were fixed in 4% paraformaldehyde (PFA) for 1 h and collected on a FACSCalibur (BD Biosciences, San Jose, CA). Analysis was performed on CellQuest software (BD Pharmingen) or FlowJo software (Tree Star, Ashland, OR).

Single-cell suspensions of the lung were prepared as described above. The lung cells were then cultured at 0.5 × 106 cells/ml in the presence of 2 μM monensin and with or without anti-CD3 (0.1 μg/ml) and anti-CD28 (1 μg/ml) for 4 h (37°C and 5% CO2) in 24-well plates. Following stimulation, the cells were collected, spun down, and stained for T lymphocyte surface markers as described above; following fixation with 100 μl of 4% PFA, the cells were transferred to a 96-well plate, washed two times with permeabilization buffer (FACS buffer plus 0.1% saponin), stained with anti-IFN-γ (2 μl, neat; BD Pharmingen) for 20 min at 4°C, and then washed two times with permeabilization buffer and once with FACS buffer. The cells were then resuspended in 4% PFA and collected on the FACSCalibur (BD Biosciences).

Tissue samples for histological studies were fixed in 10% normal buffered formalin followed by paraffin embedment. For histopathological studies, 5- to 6-μm sections were stained with Harris’ hematoxylin (Sigma-Aldrich).

Five- to 6-μm sections of formalin-fixed, paraffin-embedded lung were stained for CCL5, using anti-CCL5 (RANTES) Ab (Santa Cruz Biotechnology, Santa Cruz, CA). Immunohistochemistry was performed as recommended by manufacturer, using the goat ABC (avidin/biotin) Staining System (Vector Laborabories, Burlingame, CA). Briefly, the slides were deparaffinized and hydrated using a xylene, EtOH, and water gradient. Ag unmasking was performed with citrate buffer by microwaving the slides twice for 5 min. Following a wash in PBS and blocking with 2% donkey serum, the slides were stained with anti-CCL5 or goat anti-mouse IgG (at 1:50 in 2% donkey serum) overnight at 4°C. The slides were washed, and the secondary donkey anti-goat IgG- biotin (1:200) was applied for 30 min at room temperature. Following additional washes, the sections were incubated with avidin-biotin enzyme reagent for 30 min. Following additional washes, the sections were developed in peroxidase substrate (3-amino-9-ethylcarbazole substrate with 20 ml of N,N-dimethlyformamide in 0.02 M Na acetate buffer plus 200 μl of H2O2) for 4 min. The sections were then washed with deionized H2O and counterstained by dipping aqueous hematoxylin. After rinsing the sections, they were brightened in PBS and air-dried. Crystal mount was applied overnight, and then the sections were coverslipped.

TUNEL staining was performed on formalin-fixed, paraffin-embedded lung sections. The procedure was followed exactly as described by the ApopTag Manual using the ApopTag Peroxidase In Situ Apoptosis Detection kit (Serologicals, Norcross, GA).

Apoptosis staining on lung cells was also performed using annexin V and 7-aminoactinomycin D (7-AAD) in conjunction with surface staining for lymphocytic markers. In short, after surface staining as described above, 1 × 106 cells were washed in 100 μl of binding buffer (BD Pharmingen), resuspended in 50 μl of binding buffer, and stained with annexin V and 7-AAD (3 μl each, neat) for 15 min at room temperature. The cells were then washed with binding buffer and fixed in a 1:2 solution of 4% PFA:binding buffer. Cells were collected on the FACSCalibur within 1 h of staining.

A multiprobe RPA system (BD Pharmingen) was used to determine the levels of mRNA for genes of interest at 10-day intervals post-aerosol infection in murine lung or infected macrophages in vitro. At the time of harvest, the lungs were snap frozen in liquid nitrogen and stored at −80°C. Total RNA was extracted using TRIzol reagent (Invitrogen Life Technologies, Carlsbad, CA) followed by treatment with RNase-free DNase (Roche, Indianapolis, IN) and RNase inhibitor (Roche). Macrophage RNA was obtained from bone marrow-derived macrophages by treating ∼4 × 106 adherent cells with 1 ml of TRIzol reagent. The RNA was subjected to RPA according to BD Pharmingen’s protocol. In short, mRNA from macrophages or the lungs was hybridized overnight to [32P]UTP-labeled probes. The protected [32P]UTP-labeled RNA probes were resolved on a 6% polyacrylamide gel and analyzed by autoradiography. Cytokine analysis was performed using a custom-made template set specific for NO synthase (NOS)2, IL-4, IL-12p40, TNF-α, IL-1β, IL-1α, and IFN-γ. Chemokine analysis was performed using mCK5 multiprobe template set (BD Pharmingen). The expression of specific genes was quantified on a densitometer (ImageQuant software; Molecular Dynamics, Sunnyvale, CA), relative to the abundance of the housekeeping gene L32. We also performed phosphorimaging quantification on some samples but found results identical with densitometry; therefore, only densitometry quantifications are shown.

RNA was isolated from the lung using the TRIzol isolation protocol with slight modifications. The lung was homogenized in 3 ml of TRIzol reagent, and then two chloroform extractions were performed. Following an isopropanol precipitation, the RNA was washed with 70% ethanol and treated with RNase inhibitor (Applied Biosystems, Foster City, CA) for 45 min. Following treatment at 65°C for 15 min, the RNA was cleaned and DNase digested using the Qiagen RNA isolation kit, as directed by the manufacturer (Qiagen, Valencia, CA). The RNA was reverse transcribed using Superscript II enzyme, as directed by the manufacturer (Invitrogen Life Technologies). For real-time RT-PCR, we used the relative gene expression method (28). Hypoxanthine phosphoribosyltransferase was used as the normalizer, and uninfected lung or macrophages served as the calibrator. Each primer and probe set was tested for efficiency (all primer/probe sets had efficiencies of >97%). All samples were run in triplicate and with no-reverse-transcriptase controls on an Applied Biosystems Prism Sequence Detector 7700. Relative gene expression was calculated as 2−ΔΔCt, where ΔCt = Ctgene of interest − Ctnormalizer, and ΔΔCt = ΔCtsample − ΔCtcalibrator. Results are expressed as relative gene expression to uninfected samples. The primer and probe concentrations were used as suggested by Applied Biosystems, with the final concentration of each primer at 400 nM and probe at 250 nM.

The ELISPOT plates (96-well filtration plates; Millipore MultiScreen MAIPS4510; Millipore, Bedford, MA) were pretreated with 95% EtOH and then washed three times with sterile PBS. The rat anti-IFN-γ capture Ab (clone R4-6A2; BD Pharmingen) was added to the wells overnight at a concentration of 10 μg/ml. The ELISPOT plate was washed (all washes, PBS/0.1% Tween 20), and the plate was blocked with T cell medium containing 20% FBS for 2 h. Single-cell suspensions were prepared as described above. A total of 160,000 lymph node cells per well was incubated for 40–48 h in the presence of 20 U/ml IL-2 and with medium alone, uninfected bone marrow-derived DCs, M. tuberculosis-infected DCs (MOI, 3–4), or 10 μg/ml Con A. Following the incubation at 37°C in 5% CO2 for 48 h, the cells were washed off, and the biotinylated anti-IFN-γ detection Ab (clone XMG1.2; BD Pharmingen) was added for 2 h at 37°C at 5 μg/ml, followed by more washes and the addition of streptavidin-peroxidase for 1 h at room temperature in a humidified chamber (Vectastain ABC kit; Vector Laboratories). After additional washing, the plate was developed using the 3-amino-9-ethylcarbazole substrate kit as directed by the manufacturer. ELISPOT plates were read on the Immunospot CTL plate reader (Cellular Technology, Cleveland, OH).

Three to four mice per group per time point were used for all of the studies. Statistical analysis was performed on the data using Prism software for an unpaired t test. For bacterial numbers and cell numbers, log transformation was performed before statistical analysis to normalize the data.

CCR5 and its ligands are associated with the migration of Th1 cells and macrophages, two cell types important in a protective immune response to M. tuberculosis. Expression of these ligands was addressed during in vitro and in vivo M. tuberculosis infection. Infection of bone marrow-derived macrophages with M. tuberculosis induced CCL3, CCL4, and CCL5 expression (Fig. 1,A). RNA from the lungs of C57BL/6 mice infected with M. tuberculosis via aerosol was isolated, and real-time RT-PCR was performed on the RNA (Fig. 1,B). The expression of all known CCR5 ligands increased within the first 2 wk of infection. CCL4 had the lowest relative expression of the CCR5 ligands. The expression of CCL5, when compared with other CC chemokines was very high. Through immunohistochemical staining for CCL5 on granulomatous lung sections, we observed CCL5 protein expressed locally in both macrophage and lymphocytic areas of the granuloma (Fig. 1 C). CCR5 mRNA levels also increased in the mice following infection (data not shown). Whether this is an up-regulation of CCR5 or merely a reflection of cells expressing CCR5 migrating into the lungs cannot be determined by this assay, and an appropriate anti-CCR5 Ab for use in flow cytometry is not available.

To determine whether CCR5 plays a role in control of M. tuberculosis infection, B6;129PF2/J, C57BL/6, and CCR5−/− mice were infected via the aerosol route with ∼50 CFU. The survival of CCR5−/− mice was unaffected by M. tuberculosis infection. The mice survived >180 days, at which point experiments were terminated. There was no difference in bacterial loads, as determined by CFU, in the lungs (Fig. 2,A), liver, or spleen (data not shown). Intravenous infection of CCR5−/− mice with 2 × 105 CFU delivered via the tail vein, showed similar results (Fig. 2,B). In contrast, by 9 wk postaerosol infection, there were significantly higher bacterial loads in the lymph nodes of CCR5−/− mice (Fig. 2 C).

Because CCR5 plays a role in migration of T cells and macrophages, we examined the cellular infiltrate in the lungs following infection. At predetermined time points, the lungs were removed, the total number of cells was calculated, and flow cytometric analysis was performed to determine cell populations. The single-cell suspension was stained for T cell markers (CD4, CD8, and activation marker CD69) and macrophage and neutrophil differentiating markers (CD11b and Gr1). During initial infection, the number of cells in the lungs was not significantly different between CCR5−/− mice and wild-type mice. However, by 9 wk postinfection, the numbers of CD4 and CD8 lymphocytes were significantly higher in CCR5−/− mice (Fig. 3). The number of macrophages and neutrophils was also significantly greater in CCR5−/− than wild-type mice at 9 wk postinfection. The increased lymphocytic infiltrate in CCR5−/− mice was still evident at 6 mo postinfection (Table I). The CFU in the lungs were not statistically different at this or any time point between CCR5−/− and wild-type mice (Table I).

The engagement of CCR5, especially by CCL5, plays a role in T lymphocyte activation in some infections (29). Therefore, the expression of early activation marker CD69 on the lymphocytes in the lungs was examined. There was no difference in the expression of CD69 on CD4+ or CD8+ lymphocytes, and the percentage of CD69+ lymphocytes was similar throughout the course of infection (data not shown).

Histological analysis of the lung revealed that organized granulomas did form in CCR5−/− mice. These granulomas were similar to wild-type mice granulomas at early time points. By 9 wk postinfection, the increased lymphocytic infiltrate in the lungs of CCR5−/− mice compared with lungs of wild-type mice demonstrated by flow cytometry was evident in the histologic sections as well (Fig. 4).

The results above indicated that CCR5−/− mice had no deficiency in cell migration to the lung, but rather had more cells migrating to the lung in response to infection. To address potential mechanisms that could account for increased cell migration in the absence of CCR5, we looked for increased expression of CCR5 ligands and their alternative receptors, CCR1 and CCR3. Real-time RT-PCR was used to determine whether there was increased expression of CCL3, CCL4, or CCL5 in the lungs of CCR5−/− mice (data not shown). There was significantly greater CCL3 expression in CCR5−/− mice by day 41 post-aerosol infection (p = 0.03); although there was a trend toward greater expression of other inflammatory chemokines, such as CXCL10 and CCL2, these differences were not significant. At 42 days postinfection in the lymph nodes, the expression of CCL3, CCL4, CCL5, CXCL9, and CXCL10 was similar between CCR5−/− and wild-type mice (data not shown). Through RPAs on lung RNA, no difference in the expression of CCR1 or CCR3 was detected in the CCR5−/− or wild-type mice.

Control of M. tuberculosis infection depends on the activation of macrophages by IFN-γ and TNF, leading to the induction of inducible NOS (NOS2) and production of reactive nitrogen intermediates, such as NO. To determine whether these components of the immune response were affected by CCR5 deficiency, RPA was performed on total lung RNA (Fig. 5,A). Early after infection, the expression of IL-12, TNF, IFN-γ, and NOS2 was similar in CCR5−/− and wild-type mice, but by day 36, the CCR5−/− mice had higher expression of these genes. These were significant differences in expression of NOS2 (Fig. 5,A) and TNF (data not shown) at day 36, but variability at day 45 resulted in differences that did not reach statistical significance. Relative units in the RPA is standardized to L32, a housekeeping gene; therefore, the expression is relative on a per-cell basis. Even though there were not significant differences per cell at later time points, because there were significantly more lymphocytes in the CCR5−/− mice, there was clearly higher expression of lymphocytic genes, such as IFN-γ and TNF, overall. The IFN-γ gene expression differences were verified by intracellular cytokine staining (Fig. 5 B), and through these studies, it is evident that the increased expression of IFN-γ was due to an increase in the percentage of CD8+IFN-γ+ lymphocytes in the lungs of CCR5−/− mice.

There were a number of possible explanations for the higher numbers of lymphocytes in the lungs of CCR5−/− mice in response to M. tuberculosis infection. We hypothesized that there was less apoptosis in the CCR5−/− mice; therefore, the cells were accumulating in the lungs instead of undergoing normal turnover. Using both TUNEL staining for apoptotic bodies and flow cytometric analysis for 7-AAD and annexin V, there was no difference in the percentage of apoptotic cells between the CCR5−/− mice and wild-type mice (data not shown), but due to the increase in lymphocytes present in the lungs, there were actually significantly more apoptotic lymphocytes in the CCR5−/− mice.

We also hypothesized that CCR5 was required to recruit regulatory T lymphocytes to the lungs during M. tuberculosis infection in mice, and that this absence could interfere with normal immune regulation in the lungs. Using CD4+CD25+CD45RBlow as phenotypic markers for a regulatory T cell population (30), flow cytometric analysis of lung cells demonstrated that significantly greater numbers of these cells infiltrated the CCR5−/− mice by 6 wk postinfection, indicating that CCR5 was not required for their recruitment to the lung (data not shown).

The lung-draining lymph nodes of CCR5−/− mice contained more M. tuberculosis bacilli than the lymph nodes of wild-type mice (Fig. 2 C). We hypothesized that the increased bacterial load was due to greater numbers of DCs migrating from the lungs to the lymph nodes carrying M. tuberculosis. As a result, more T lymphocytes were being primed in the lymph node and then migrating to the lungs. To explore this hypothesis, flow cytometric analysis was performed on the mediastinal lymph nodes of CCR5−/− mice following M. tuberculosis infection.

The overall total cell numbers of the lymph nodes of CCR5−/− and B6;129PF2/J mice were similar, but at 6 and 9 wk postinfection,there were statistically significant differences in the total number of DCs (defined as CD11c+CD11bhigh (or low)) in the CCR5−/− mice (Fig. 6,A). ELISPOT analysis confirmed that significantly more lymphocytes from lymph nodes of CCR5−/− mice were primed to produce IFN-γ when stimulated by either Con A or M. tuberculosis-infected DCs (Fig. 6 B).

In CCR2−/− mice, the requirement for CCR2 in control of M. tuberculosis infection was dose dependent. These mice were unable to control M. tuberculosis when it is administered at a standard dose of 2.5 × 105 CFU delivered i.v., but could control lower dose aerosol or i.v. infection (31). We investigated whether the ability of CCR5−/− mice to control infection was route or dose dependent. CCR5−/− mice infected i.v. (2 × 105 CFU/mouse) controlled M. tuberculosis growth, and the cell migration pattern was similar to that of CCR5−/− mice infected via the aerosol route (Fig. 1 B). The CCR5−/− mice again had strikingly high cell infiltration to their lungs, but with no apparent long-term detrimental effects of this increased infiltration.

CCR5 is expressed on many immune cells in the lung following M. tuberculosis aerosol infection in mice, and the ligands for this receptor are induced in the lungs by this infection. Whether CCR5 is required for the migration of these cells to the lungs and subsequent control of infection was unknown. The results of this study provide evidence that CCR5 is not required for migration of immune cells to the lungs or for the initiation of the Th1 immune response during M. tuberculosis infection, but may influence the pathology directly or indirectly. CCR5 may affect the retention of DCs in the lungs and migration to the lymph nodes. CCR5−/− mice had more bacilli in the lung-draining lymph nodes, as well as more DCs. This coincided with enhanced priming of M. tuberculosis-specific T cells in the lymph nodes. Such increased priming likely accounts for the substantially increased numbers of T lymphocytes migrating to the lungs of CCR5−/− mice following M. tuberculosis infection. It is also possible that the lack of CCR5 prevents immune control of M. tuberculosis replication in the lymph nodes, accounting for more bacteria at this site. However, there is no apparent dependence on CCR5 for control of M. tuberculosis in the lungs.

CCR5 function may be compensated for by other chemokine receptors and chemokines expressed in the lungs during infection. Although CCL4 and CCL5 can also signal through CCR1, the immune cells express many other chemokine receptors that may be responsible for the migration of these cells to the lung. These include CXCR3, CCR2, CCR1, and CXCR2. There is no reported increase in susceptibility to tuberculosis in humans carrying a deletion in the CCR5 gene (CCR5Δ32).

The finding that CCR5−/− mice had more inflammation in their lungs was intriguing and unexpected. Recent data in other systems support an immunoregulatory role for CCR5. CCR5−/− mice had enhanced delayed-type hypersensitivity and humoral responses to T cell-dependent antigenic challenge (23). When infected with influenza A virus, CCR5−/− mice had a hyperinflammatory response, resulting in increased pulmonary inflammation and a decrease in survival (24). In a murine model of colitis using dextran sodium sulfate, CCR5-deficient mice had increased infiltration of CD4+ and NK1.1+ lymphocytes, along with a decrease in Th1 and an increase in Th2 cytokine expression (32). Unfortunately, none of these studies have been able to define a mechanism for the heightened immune response in the CCR5−/− mice.

We explored several possibilities for the increased numbers of immune cells in the lungs of the M. tuberculosis-infected CCR5−/− mice, in the absence of increased bacterial numbers. One hypothesis was that there was a decrease in cell turnover in the lungs of CCR5−/− mice. Recent studies have suggested that CCR5 signaling can induce apoptosis. The HIV protein Env can activate CCR5 to induce the Fas and caspase-8 pathway resulting in CD4+ cell death (33). However, in our studies, there were equal or greater numbers of apoptotic cells in the CCR5−/− mice as detected by two different methods, discounting this hypothesis in this system.

We also assessed whether CCR5 was important for migration of regulatory T lymphocytes to the lungs and the down-regulation of Th1 responses, which could account for the observed increased inflammation in the absence of CCR5. Using the best markers known for regulatory T cells (30), there were significantly more cells of this phenotype in the lungs of CCR5−/− mice. Therefore, CCR5 does not appear to be playing a role in regulatory T cell migration, but rather, the increased migration may be a response to the increased inflammation, or simply related to the overall increase in cell numbers in the lungs of these mice.

An alternative hypothesis was that changes in the concentration of bound chemokines can lead to increased cell migration to the lungs. Although there is a deletion of CCR5 in these mice, CCL3, CCL4, and CCL5 were still being produced, and therefore, the concentration of unbound chemokine may be higher in the CCR5−/− mice. In addition, there was increased mRNA expression of some chemokines (such as CCL3) in the lungs of the CCR5−/− mice compared with wild-type mice. Such increased expression/circulation of chemokines may lead to signaling through receptors other than CCR5 in the lungs and could result in the influx of immune cells. However, the increased expression may also have been a reaction to the numbers of cells in the lungs.

A final hypothesis was that CCR5 is required for normal DC migration. We observed greater bacterial numbers in the mediastinal lymph nodes of CCR5−/− mice. DCs infected in the lungs are likely migrating and carrying M. tuberculosis bacilli to the lymph nodes. Previous studies demonstrated maturation of DCs upon infection with M. tuberculosis (5, 10, 34, 35); and mature DCs migrate to the lymph nodes (36, 37, 38) where they prime naive T cells. At 6 and 9 wk postinfection, there were more DCs in the lymph nodes of CCR5−/− mice. The increased Ag dose (i.e., bacterial numbers) in conjunction with increased numbers of DCs within the lymph nodes likely leads to enhanced priming of lymphocytes, as measured in the ELISPOT assays. More primed T lymphocytes then are available to migrate to the site of infection in the lungs, resulting in the higher T lymphocyte numbers observed in the CCR5−/− mice.

Upon maturation, DCs are believed to down-regulate expression of CCR5 and CCR6, whereas they up-regulate other receptors such as CXCR4 and CCR7, targeting these cells to the lymph nodes (reviewed in Ref. 39). We observed increased numbers of DCs in the lymph nodes following infection in CCR5−/− mice, suggesting that, during M. tuberculosis infection, the expression of β chemokines such as CCL3, CCL4, and CCL5 may normally retain CCR5+ DCs in the lungs or recruit the DCs out of the lymph nodes to the lungs. The absence of CCR5 apparently results in either increased migration from the lungs to the lymph nodes, or retention of DCs in the lymph nodes.

This study supports our previous finding that, although there may not be striking differences in bacterial loads in the lungs of some transgenic mouse models, these models are very useful for studying the immune response (14). Addressing the function of molecules in the immune response requires careful experimental design and execution, and interpretation of experimental results. Roles may exist for molecules in the immune response even if the mice are controlling infection at the level of bacterial burden. The immune response in the C57BL/6 and B6;129PF2/J mice, although not sufficient to eliminate bacterial numbers in the lungs, does control the infection. In this study, as well as other models (IL-10−/−, for example; H. M. Scott Algood, J. Chan, and J. L. Flynn, unpublished data), enhancing the type 1 T cell response or the ability of T lymphocytes to enter the lungs over the wild-type level, does not have an obvious beneficial effect on control of the infection. These data indicate that there are additional factors that must be induced or enhanced to increase the ability of the host to eliminate M. tuberculosis infection.

In summary, our study suggests a role for CCR5 in immune regulation. CCR5 may be controlling DC migration to the lymph nodes, possibly by maintaining these cells in infected lungs. Thus, the absence of CCR5 resulted in increased numbers of both DCs and M. tuberculosis bacilli in lymph nodes; we assume that the bacteria were carried from the lungs in DCs. The result of the increased numbers of DCs and Ags was enhanced T lymphocyte priming, leading to increased inflammation in the lungs. There may also be increased levels of unbound chemokine inducing migration of immune cells through receptors other than CCR5, increasing the overall cellular infiltrate and size of the granulomas.

We are grateful to Amy Myers for excellent technical assistance. We thank Dr. John Chan and the members of the Flynn and Nau laboratories for helpful discussions.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by National Institutes of Health Grant HL71241 (to J.L.F.), American Lung Association Grant CI-016-N, Western Pennsylvania Lung Association Dissertation Grant (to H.M.S.A.), and National Institutes of Health T32 AI49820 (to H.M.S.A.).

3

Abbreviations used in this paper: DC, dendritic cell; MOI, multiplicity of infection; PFA, paraformaldehyde; 7-AAD, 7-aminoactinomycin D; RPA, RNase protection assay; NOS, NO synthase; Ct, cycle threshold.

1
World Health Organization.
2003
.
Global Tuberculosis Control: Surveillance, Planning, Financing: WHO Report 2003
World Health Organization, Geneva, Switzerland.
2
Flynn, J. L., J. Chan.
2001
. Immunology of tuberculosis.
Annu. Rev. Immunol.
19
:
93
.
3
Gonzalez-Juarrero, M., O. C. Turner, J. Turner, P. Marietta, J. V. Brooks, I. M. Orme.
2001
. Temporal and spatial arrangement of lymphocytes within lung granulomas induced by aerosol infection with Mycobacterium tuberculosis.
Infect. Immun.
69
:
1722
.
4
Uehira, K., R. Amakawa, T. Ito, K. Tajima, S. Naitoh, Y. Ozaki, T. Shimizu, K. Yamaguchi, Y. Uemura, H. Kitajima, et al
2002
. Dendritic cells are decreased in blood and accumulated in granuloma in tuberculosis.
Clin. Immunol.
105
:
296
.
5
Gonzalez-Juarrero, M., I. M. Orme.
2001
. Characterization of murine lung dendritic cells infected with Mycobacterium tuberculosis.
Infect. Immun.
69
:
1127
.
6
Feng, C. G., A. G. Bean, H. Hooi, H. Briscoe, W. J. Britton.
1999
. Increase in γ-interferon-secreting CD8+, as well as CD4+, T cells in lungs following aerosol infection with Mycobacterium tuberculosis.
Infect. Immun.
67
:
3242
.
7
Serbina, N. V., J. L. Flynn.
1999
. Early emergence of CD8+ T cells primed for production of type 1 cytokines in the lungs of Mycobacterium tuberculosis-infected mice.
Infect. Immun.
67
:
3980
.
8
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
.
9
Chan, J., Y. Xing, R. Magliozzo, B. R. Bloom.
1992
. Killing of virulent Mycobacterium tuberculosis by reactive nitrogen intermediates produced by activated murine macrophages.
J. Exp. Med.
175
:
1111
.
10
Bodnar, K. A., N. V. Serbina, J. L. Flynn.
2001
. Fate of Mycobacterium tuberculosis within murine dendritic cells.
Infect. Immun.
69
:
800
.
11
Tailleux, L., O. Neyrolles, S. Honore-Bouakline, E. Perret, F. Sanchez, J. P. Abastado, P. H. Lagrange, J. C. Gluckman, M. Rosenzwajg, J. L. Herrmann.
2003
. Constrained intracellular survival of Mycobacterium tuberculosis in human dendritic cells.
J. Immunol.
170
:
1939
.
12
Lazarevic, V., J. Flynn.
2002
. CD8+ T cells in tuberculosis.
Am. J. Respir. Crit. Care Med.
166
:
1116
.
13
Mack, M., J. Cihak, C. Simonis, B. Luckow, A. E. Proudfoot, J. Plachy, H. Bruhl, M. Frink, H. J. Anders, V. Vielhauer, et al
2001
. Expression and characterization of the chemokine receptors CCR2 and CCR5 in mice.
J. Immunol.
166
:
4697
.
14
Scott Algood, H. M., J. Chan, J. F. Flynn.
2003
. Chemokines and tuberculosis.
Cytokine Growth Factor Rev.
14
:
467
.
15
Scott Algood, H. M., J. L. Flynn.
2004
. TNF influences chemokine expression of macrophages in vitro and CD11b+ cells in vivo during Mycobacterium tuberculosis infection.
J. Immunol.
172
:
6846
.
16
Sadek, M. I., E. Sada, Z. Toossi, S. K. Schwander, E. A. Rich.
1998
. Chemokines induced by infection of mononuclear phagocytes with mycobacteria and present in lung alveoli during active pulmonary tuberculosis.
Am. J. Respir. Cell Mol. Biol.
19
:
513
.
17
Rhoades, E. R., A. M. Cooper, I. M. Orme.
1995
. Chemokine response in mice infected with Mycobacterium tuberculosis.
Infect. Immun.
63
:
3871
.
18
Wu, L., G. LaRosa, N. Kassam, C. J. Gordon, H. Heath, N. Ruffing, H. Chen, J. Humblias, M. Samson, M. Parmentier, et al
1997
. Interaction of chemokine receptor CCR5 with its ligands: multiple domains for HIV-1 gp120 binding and a single domain for chemokine binding.
J. Exp. Med.
186
:
1373
.
19
Fraziano, M., G. Cappelli, M. Santucci, F. Mariani, M. Amicosante, M. Casarini, S. Giosue, A. Bisetti, V. Colizzi.
1999
. Expression of CCR5 is increased in human monocyte-derived macrophages and alveolar macrophages in the course of in vivo and in vitro Mycobacterium tuberculosis infection.
AIDS Res. Hum. Retroviruses
15
:
869
.
20
Sato, N., W. A. Kuziel, P. C. Melby, R. L. Reddick, V. Kostecki, W. Zhao, N. Maeda, S. K. Ahuja, S. S. Ahuja.
1999
. Defects in the generation of IFN-γ are overcome to control infection with Leishmania donovani in CC chemokine receptor (CCR)5-, macrophage inflammatory protein-1α-, or CCR2-deficient mice.
J. Immunol.
163
:
5519
.
21
Huffnagle, G. B., L. K. McNeil, R. A. Mcdonald, J. W. Murphy, G. B. Toews, N. Maeda, W. A. Kuziel.
1999
. Cutting edge: role of C-C chemokine receptor 5 in organ-specific and innate immunity to Cryptococcus neoformans.
J. Immunol.
163
:
4642
.
22
Aliberti, J., C. Reis e Sousa, M. Schito, S. Hieny, T. Wells, G. B. Huffnagle, A. Sher.
2000
. CCR5 provides a signal for microbial induced production of IL-12 by CD8α+ dendritic cells.
Nat. Immunol.
1
:
83
.
23
Zhou, Y., T. Kurhara, R. P. Ryseck, Y. Yang, C. Ryan, J. Loy, G. Warr.
1998
. Impaired macrophage function an enhanced T cell-dependent immune response in mice lacking CCR5, the mouse homologue of the major HIV-1 coreceptor.
J. Immunol.
160
:
4018
.
24
Dawson, T. C., M. A. Beck, W. A. Kuziel, F. Henderson, N. Maeda.
2000
. Contrasting effects of CCR5 and CCR2 deficiency in the pulmonary inflammatory response to influenza A virus.
Am. J. Pathol.
156
:
1951
.
25
Serbina, N. V., C. C. Liu, C. A. Scanga, J. L. Flynn.
2000
. CD8+ CTL from lungs of Mycobacterium tuberculosis-infected mice express perforin in vivo and lyse infected macrophages.
J. Immunol.
165
:
353
.
26
Flynn, J. L., M. M. Goldstein, K. J. Triebold, B. Koller, B. R. Bloom.
1992
. Major histocompatibility complex class I-restricted T cells are required for resistance to Mycobacterium tuberculosis infection.
Proc. Natl. Acad. Sci. USA
89
:
12013
.
27
Olszewski, M. A., G. B. Huffnagle, T. R. Traynor, R. A. McDonald, D. N. Cook, G. B. Toews.
2001
. Regulatory effects of macrophage inflammatory protein 1α/CCL3 on the development of immunity to Cryptococcus neoformans depend on expression of early inflammatory cytokines.
Infect. Immun.
69
:
6256
.
28
Liu, W., D. A. Saint.
2002
. A new quantitative method of real time reverse transcription polymerase chain reaction assay based on simulation of polymerase chain reaction kinetics.
Anal. Biochem.
302
:
52
.
29
Luther, S. A., J. G. Cyster.
2001
. Chemokines as regulators of T cell differentiation.
Nat. Immunol.
2
:
102
.
30
Shevach, E. M..
2002
. CD4+CD25+ suppressor T cells: more questions than answers.
Nat. Rev. Immunol.
2
:
389
.
31
Scott, H. M., J. L. Flynn.
2002
. Mycobacterium tuberculosis in chemokine receptor 2-deficient mice: influence of dose on disease progression.
Infect. Immun.
70
:
5946
.
32
Andres, P. G., P. L. Beck, E. Mizoguchi, A. Mizoguchi, A. K. Bhan, T. Dawson, W. A. Kuziel, N. Maeda, R. P. MacDermott, D. K. Podolsky, H. C. Reinecker.
2000
. Mice with a selective deletion of the CC chemokine receptors 5 or 2 are protected from dextran sodium sulfate-mediated colitis: lack of CC chemokine receptor 5 expression results in a NK1.1+ lymphocyte-associated Th2-type immune response in the intestine.
J. Immunol.
164
:
6303
.
33
Algeciras-Schimnich, A., S. R. Vlahakis, A. Villasis-Keever, T. Gomez, C. J. Heppelmann, G. Bou, C. V. Paya.
2002
. CCR5 mediates Fas- and caspase-8 dependent apoptosis of both uninfected and HIV-infected primary human CD4 T cells.
AIDS
16
:
1467
.
34
Henderson, R. A., S. C. Watkins, J. L. Flynn.
1997
. Activation of human dendritic cells following infection with Mycobacterium tuberculosis.
J. Immunol.
159
:
635
.
35
Tascon, R. E., C. S. Soares, S. Ragno, E. Stavropoulos, E. M. Hirst, M. J. Colston.
2000
. Mycobacterium tuberculosis-activated dendritic cells induce protective immunity in mice.
Immunology
99
:
473
.
36
Legge, K. L., T. J. Braciale.
2003
. Accelerated migration of respiratory dendritic cells to the regional lymph nodes is limited to the early phase of pulmonary infection.
Immunity
18
:
265
.
37
Bozza, S., R. Gaziano, A. Spreca, A. Bacci, C. Montagnoli, P. di Francesco, L. Romani.
2002
. Dendritic cells transport conidia and hyphae of Aspergillus fumigatus from the airways to the draining lymph nodes and initiate disparate Th responses to the fungus.
J. Immunol.
168
:
1362
.
38
Vermaelen, K. Y., I. Carro-Muino, B. N. Lambrecht, R. A. Pauwels.
2001
. Specific migratory dendritic cells rapidly transport antigen from the airways to the thoracic lymph nodes.
J. Exp. Med.
193
:
51
.
39
Caux, C., B. Vanbervliet, C. Massacrier, S. Ait-Yahia, C. Vaure, K. Chemin, M.-C. Dieu-Nosjean, A. Vicari.
2002
. Regulation of dendritic cell recruitment by chemokines.
Transplantation
73
:
S7
.