Hematopoietic Cells Are Required to Initiate a Chlamydia trachomatis-Specific CD8⁺ T Cell Response

Mice

Female BALB/cBy/J mice (H-2^d), female C57BL/6 mice (H-2^b), and male CbyB6F₁/J mice (H-2^b/d) were purchased from The Jackson Laboratory (Bar Harbor, ME). Mice were housed in a pathogen-free barrier facility at Harvard Medical School and used at 8–12 wk of age, unless otherwise indicated.

Tissue culture

BALB/3T3 (H-2^d fibroblasts from a BALB/c embryo), P815 (an H-2^d mastocytoma from a DBA/2 mouse), J774 (H-2^d Mφ from a BALB/c mouse), and 1308.1 (an H-2^b thymic stromal cell line) cells were grown at 37°C in 7.0% CO₂ in a medium (RP10) that consisted of RPMI 1640 supplemented with 2 mM l-glutamine, 50 μM 2-ME, antibiotics (when indicated), 5 mM HEPES, and 10% FCS.

Growth, isolation, and detection of C. trachomatis

EBs of C. trachomatis serovar L2 434/Bu were propagated within McCoy cell monolayers grown in Eagle’s MEM supplemented with 1.5 g/L sodium bicarbonate, 0.1 mM nonessential amino acids, 1 mM sodium pyruvate, and 10% FCS. EBs were released from infected McCoy cells with glass beads and purified by density gradient centrifugation, as described (32). Aliquots of C. trachomatis were stored at −80°C in a buffer containing 250 mM sucrose, 10 mM sodium phosphate, and 5 mM l-glutamic acid (pH 7.2) (sucrose-phosphate-glutamate freezing media) (33).

To measure the productivity of C. trachomatis infection in BALB 3T3 cells, bone marrow-derived Mφs, and bone marrow-derived DCs, cells were infected for the indicated periods of time and then frozen at −80°C. Cells were subsequently thawed and sonicated to release viable EBs. Lysates of infected cells were used to infect McCoy cell monolayers and incubated for 36 h to allow the development of inclusions. McCoy cell monolayers were infected in duplicate for each time point. Monolayers were then fixed and stained with a FITC-conjugated anti-C. trachomatis Ab to visualize developing inclusions. For each well, three microscopic fields were chosen at random and the number of inclusion-forming units (IFU)/field was counted. The mean number of IFU/field was then calculated by averaging the number of IFU/field across duplicate wells. To determine the mean number of IFU/sample, the following calculation was made: mean number of IFU/sample = mean number of IFU/field × lens factor* × dilution factor**. *, Refers to the fraction of the total area of the cell monolayer that is viewed under a particular objective. **, Refers to the fraction of the original sample that actually gets plated onto McCoy cell monolayer. The productivity of infection of each cell type was reported as the ratio of the mean number of IFU recovered from host cells at a given time point (i.e., output IFU) relative to the average number of IFU used to initially infect host cells (i.e., input IFU).

Infection of tissue culture monolayers with C. trachomatis

Before infection, cells were rinsed twice with medium without antibiotics and incubated overnight. To infect cells with C. trachomatis, cells were incubated with bacteria in SPG and centrifuged at 1925 × g at 30°C for 1 h to increase the contact between the bacteria and the tissue culture cells. The multiplicity of infection (MOI) used for each experiment varied and is specifically mentioned in Results. After centrifugation, the inoculum was removed and the cells were incubated for the indicated periods of time at 37°C in medium without antibiotics.

Stimulation and maintenance of C. trachomatis-specific CD8⁺ T cells

The C. trachomatis-specific H-2^d-restricted CD8⁺ T cell line 69 was maintained and expanded by weekly stimulation on C. trachomatis-infected J774 cells, as described (11). T cells specific for the Cap1_139–147 epitope were stimulated from infected mice, as previously described (16). Briefly, mice were infected i.p. with 1 × 10⁸ IFU of C. trachomatis serovar L2. After 14 days, splenocytes from the infected animals were washed once in RP10, and 4 × 10⁷ splenocytes were cultured with an equal number of naive, irradiated, syngeneic splenocytes that had been treated for 1 h with 10 μM Cap1_139–147 peptide. These cultures were incubated upright for 5 days in a T-25 flask at 37°C in 7% CO₂ in a total volume of 20 ml of RP10 before their use as effector cells in chromium release assays. After two rounds of restimulation, cells were subsequently restimulated in RP10 supplemented with supernatant from cultured rat splenocytes stimulated with Con A as a source of IL-2.

Assays for T cell activity

Chromium release assays were used to determine the percent specific lysis of target cells, as previously described (11). IFN-γ ELISA was also used to measure T cell activity. Supernatants from the coculture of CD8⁺ T cells and APCs were collected after 16 h, unless otherwise noted. These supernatants were analyzed for the presence of IFN-γ using an IFN-γ Minikit (Pierce, Woburn, MA), according to the manufacturer’s instructions.

IFN-γ ELISPOT

Spleens from experimental mice were harvested, and the RBC were lysed with ammonium/chloride potassium (ACK) lysis buffer (34). The splenocytes were counted and added at a concentration of 5 × 10⁵ or 1 × 10⁵ cells/well to nitrocellulose-backed 96-well plates previously coated with rat anti-mouse IFN-γ Ab (18181D; BD Biosciences, San Jose, CA). Each well also contained 1 × 10⁵ irradiated P815 stimulator cells that had been previously pulsed with Cap1_139–147 peptide. The plates were incubated for 24–28 h and then washed and treated with biotinylated rat anti-mouse IFN-γ Ab (XMG1.2; BD Biosciences). After an additional 18 h, the plates were washed again, and streptavidin-HRP (BD Biosciences) was added. The plates were developed with 1 mg/ml 3, 3′-diaminobenzidine tetrahydrochloride dyhrate (Bio-Rad, Hercules, CA) in 50 mM Tris (pH 7.9).

Isolation of bone marrow-derived Mφ and DCs

Femurs and tibias were removed from mice and flushed with RP10. After washing the bone marrow cells once, the total number of cells was counted. For the culture of Mφ, ∼1 × 10⁶ cells per plate were seeded into 100-mm nontissue culture-treated petri dishes. Mφ were cultured in RPMI 1640 medium containing 20% FCS, l-glutamine, penicillin, streptomycin, and 20% L929 cell-conditioned medium. The medium was replaced on day 4, and cells were used on day 7. DCs were cultured, as described by Lutz et al. (35). Briefly, 5 × 10⁶ bone marrow cells were cultured in RP10 supplemented with J558L cell-conditioned medium. J558L is a GM-CSF-producing mouse Mφ-like cell line that was generously provided by I. Mellman (Yale University, New Haven, CT). The medium was supplemented on day 3, and the DCs were used on day 6. The purity of the DC cultures was assessed by flow cytometry, as described below. To obtain mature DCs for use in experiments, the DCs were further incubated in culture medium containing 5 μg/ml purified LPS (Sigma-Aldrich, St. Louis, MO) for 16 h.

Paraformaldehyde fixation of APCs

For experiments in which APCs were fixed before their use in assays, the cells were distributed into 96-well plates. After infection and incubation, cells were centrifuged for 10 min at 260 × g and washed once with RPMI 1640. Cells were fixed with a 1% paraformaldehyde solution for 10–15 min at room temperature. The cells were then washed twice in RPMI 1640 before their use in assays.

Flow cytometric analysis of DC culture purity and maturity

Cells were surface stained and analyzed using standard protocols (36). Abs specific for the following molecules were obtained from BD Biosciences: DC/Mφ markers, CD11b (M1/70) and CD11c (HL3); DC activation markers, CD86 (GL1), CD80 (16-10A1), and CD40 (HM40-3); and a MHC class II molecule: I-A^b (AF6-120.1) or I-A^d (39-10-8). Cells were stained, washed, and resuspended in HBSS containing 1.0% BSA and 0.1% NaN₃.

Magnetic cell separation of DCs

For adoptive transfer studies, DCs were purified using the MACS magnetic bead system, according to the manufacturer’s instructions (Biotech International, Auburn, CA). Cells were washed and resuspended in PBS supplemented with 0.5% BSA. Before separation, nonspecific binding of the microbeads by Fc receptors expressed by Mφ was inhibited by incubating the cells with 1 μg of Fc block reagent (BD Biosciences) per 1 × 10⁶ cells for 15 min at 4°C. After blocking, 100 μl of CD11c microbeads per 1 × 10⁸ cells was added, and the mixture was incubated for 15 min at 6–12°C. The mixture was then washed twice in PBS/0.5% BSA and resuspended in 500 μl of PBS/0.5% BSA per 1 × 10⁸ cells. Magnetic separation was initiated by application of the bead-labeled cells to a column attached to a MACS separator magnet. After washing the column, the column was removed from the magnetic field, and positively labeled cells were flushed from the column. FACS analysis of these cells using anti-CD11c and anti-CD11b Abs showed these cells to be ≥85% DCs.

Treatment of cells with chemical inhibitors of Ag processing

APCs were infected with C. trachomatis and incubated for 16–20 h to allow time for C. trachomatis CD8⁺ T cell Ags to be expressed before inhibitors were added. The following inhibitors were used: Z-L₃-VS at a concentration of 50 μM (a gift from H. Ploegh, Harvard Medical School, Boston, MA) and bafilomycin A1 at a concentration of 1 μM (Calbiochem, San Diego, CA). Both inhibitors were dissolved in DMSO and then diluted 100-fold into the cell cultures. APCs were incubated with inhibitor or DMSO alone as a negative control at 37°C for an additional 4 h. After incubation, the cells were washed thoroughly to insure the removal of both the DMSO and any unincorporated inhibitor.

Generation of radiation bone marrow chimeras

CB6 mice were irradiated with two doses of 600 rad separated by 3 h. Irradiated mice were reconstituted with 6 × 10⁶ bone marrow cells from the indicated mice on the following day. To avoid lysis of donor bone marrow cells by NK cells in the recipient, 10 μl of anti-asialo GM1 Ab (Wako Chemicals, Osaka, Japan) was injected i.p. into recipient mice on the day of bone marrow transplant and 3 days after bone marrow reconstitution. To assess the level of chimerism at the time of infection, peripheral blood cells were analyzed by flow cytometry for the presence of donor-derived cells. Mice were anesthetized with isofluorane, and their blood was collected retraorbitally. RBC were lysed with ACK lysis buffer, and the remaining white blood cells were stained with anti-TCR-β (H57-597; BD Biosciences) Abs (to evaluate the T cell phenotype) and anti-CD11b (M1/70; BD Biosciences) Abs (to evaluate the Mφ and DC phenotype). The leukocyte populations were analyzed by flow cytometry for their expression of the MHC allele of the bone marrow donor mice using anti-H-2K^d (SF1–1.1; BD Biosciences) or anti-H-2K^b (AF6-88.5; BD Biosciences) Abs. All animals used in experiments displayed 85–97% levels of chimerism, as measured by the expression of the donor cell MHC allele on peripheral blood cells from the chimeric mice. Experiments were performed from 8 wk to 6 mo after irradiation and reconstitution. All experimental groups were age matched.

Tetracycline treatment of mice

A total of 2 mg of tetracycline was administered i.p. 24 h before infection and then every 24 h until the end of the experiment (37).

In vitro priming assay

Splenocytes from naive or C. trachomatis-immune BALB/c mice were depleted of RBC using ACK lysis buffer. To enrich for T cells, leukocytes were loaded onto a mouse T cell enrichment column (R&D Systems, Minneapolis, MN). Following enrichment, the T cell populations were labeled with 5 μM CFSE, as previously described (38). Briefly, cells were resuspended in PBS/0.1% BSA at a concentration of 10⁷ cells/ml. CFSE was added to cells to a final concentration of 5 μM, and cells were labeled for 10 min at 37°C. After labeling, the cells were washed with PBS/0.1% BSA and resuspended in RP10. The labeled T cells were cocultured with the appropriate APC population at an E:T ratio of 10:1. After 3 days of in vitro stimulation, the cultures were harvested and stained with anti-CD8 (53-5.8; BD Biosciences) and anti-TCR-β (H57-597; BD Biosciences) Abs before analysis by flow cytometry.

Cross-presentation assay

The 1308.1 or bone marrow-derived Mφ or DCs from C57BL/6 mice (H-2^b) were infected with C. trachomatis and incubated for 24 h in medium without antibiotics. These cells were individually mixed at a 1:1 ratio with BALB/c (H-2^d) bone marrow-derived DCs or cultured alone in 96-well flat-bottom culture plates. The cocultures of infected H-2^b donor cells and uninfected H-2^d DCs were incubated in medium containing 1 μg/ml chloramphenicol to prevent infection of the H-2^d DCs. After 16 h of coculture, C. trachomatis-specific CD8⁺ T cell line 69 was added to the mixture of APCs at an E:T ratio of 10:1. The culture supernatant was collected after 12 h of T cell stimulation and analyzed using an IFN-γ ELISA.

C. trachomatis infection of Mφ and DCs is less productive than infection of nonprofessional APCs

It is well established that C. trachomatis replicates efficiently within nonprofessional APCs (39). To determine how the productivity of infection of Mφ and DCs compares with that of nonprofessional APCs, we infected BALB/c DCs, BALB/c Mφ, and BALB/3T3 fibroblasts cells with C. trachomatis at a MOI of 10:1. Productive C. trachomatis infection of a host cell was defined as the ability of bacteria to enter the host cell, develop into RBs, replicate, and then convert back into the infectious EB form. We assayed the ability of C. trachomatis to replicate in the three cell types by comparing the ratio of the number of EBs that exited the infected cell population for every EB that entered these cells. In all three cell types, we observed a reduction in the number of IFU recovered soon after infection (Fig. 1⇓). This was expected because at these time points the bacteria had differentiated into RBs, the noninfectious form of the organism that could not be measured by these assays. As anticipated, there was a significant increase in the number of IFU recovered from C. trachomatis-infected BALB/3T3 fibroblasts over time (Fig. 1⇓A). In contrast, the number of EBs recovered from DCs and Mφ was dramatically lower (Fig. 1⇓, B and C). Increasing the MOI to 25:1 for DCs and Mφ resulted in only minimal increases in the productivity of infection (data not shown). It is not clear whether the low productivity of infection in Mφ and DCs was simply the result of only a small number of cells being infected or the poor development of C. trachomatis-containing inclusions within a large number of infected cells. Regardless, we find that C. trachomatis infection is much more productive in nonprofessional APCs than in Mφ and DCs.

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FIGURE 1.

C. trachomatis infection of Mφ and DCs is far less productive than infection of nonprofessional APCs. BALB/3T3 (A), BALB/c DC (B), and BALB/c Mφ (C) were infected with C. trachomatis at a MOI of 10:1. At the indicated times after infection, the cells were freeze thawed and subsequently sonicated to release intracellular EBs. These lysates were then used to infect McCoy cell monolayers. Duplicate wells were infected for each sample. McCoy cell monolayers were fixed 36 h after infection and stained for inclusions using a FITC-conjugated Ab specific for C. trachomatis. The number of inclusions per microscopic field was counted for each time point, and the average IFU recovered for each sample were calculated, as described in Materials and Methods. The fold difference in the number of IFU recovered at each time point relative to the input IFU was calculated by dividing the average number of IFU per sample at the specified time point (i.e., output IFU) by the number of IFU present at t = 0 h postinfection (i.e., input IFU). Sample variation across duplicate wells never resulted in a value for the fold difference that was greater than 0.04 for DCs, 1.0 for Mφ, or 20 for BALB/3T3 cells. These differences are not significant given the levels of infection within each individual cell type. The data shown are representative of three separate experiments. The purity of both the DC and Mφ cells used in this experiment was >90%.

C. trachomatis-specific CD8⁺ T cells recognize infected Mφ and DCs

Because the low level of C. trachomatis replication in Mφ and DCs may result in undetectable levels of C. trachomatis Ag presentation by these cells, we tested whether the previously identified C. trachomatis-specific CD8⁺ T cell line 69 (11) could recognize C. trachomatis-infected Mφ and DCs (Fig. 2⇓). As expected, infected nonprofessional APCs, which support high levels of bacterial replication (Fig. 1⇑A), consistently stimulated IFN-γ secretion by the C. trachomatis-specific CD8⁺ T cell line 69 (Fig. 2⇓A). In contrast, C. trachomatis-infected Mφ, in which bacterial replication was limited (Fig. 1⇑C), stimulated ∼10-fold less IFN-γ secretion from the T cell line than did infected nonprofessional APCs (Fig. 2⇓C). Interestingly, despite our finding that DCs were the least productively infected cell type (Fig. 1⇑B), these cells stimulated the greatest amount of IFN-γ secretion from the C. trachomatis-specific CD8⁺ T cell line 69 (Fig. 2⇓B). These results show the enhanced ability of DCs to process and present Ag, even overcoming the limitation imposed by modest C. trachomatis replication.

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FIGURE 2.

C. trachomatis-specific CD8⁺ T cell line 69 recognizes infected BALB/3T3 cells, Mφ and DCs. Cells were infected with C. trachomatis for the indicated times and then fixed with paraformaldehyde. Uninfected cells were also included as targets to establish a baseline level of IFN-γ secretion in response to each cell type. CD8⁺ T cell line 69 was added at an E:T ratio of 10:1. Sixteen hours later, the amount of IFN-γ secreted into the supernatant was measured by ELISA. A, BALB/3T3 cells were infected at a MOI of 100:1. B, BALB/c DC (C) and Mφ were infected at a MOI of 50:1. Error bars represent SD of triplicate wells. The results shown are representative of three independent experiments.

Processing of C. trachomatis Ag is proteasome dependent

One aspect of Ag presentation that distinguishes Mφ and DCs from nonprofessional APCs is their ability to process both endogenous and exogenous Ags for presentation on MHC class I. The recognition of infected nonprofessional APCs by C. trachomatis-specific CD8⁺ T cells indicates that some C. trachomatis Ag accesses the host cell cytosol for processing by the endogenous MHC class I processing pathway (11). This pathway is dependent on the host cell proteasome to digest protein Ags into peptide epitopes within the host cell cytosol. However, in Mφ and DCs, Ags also can be processed through an endosomal pathway termed the exogenous MHC class I processing pathway (40). CD8⁺ T cell Ags processed within the endosomal compartment can be processed by the acid proteases that also generate the peptides that bind MHC class II molecules. We investigated whether the vacuolar localization of C. trachomatis during development might allow Ags from this pathogen to be processed by the endosomal, exogenous MHC class I processing pathway in Mφ and DCs.

To examine the relative contribution of the exogenous vs the endogenous MHC class I processing pathways to the processing of C. trachomatis Ag in infected cells, we used chemical inhibitors to selectively block the digestion of C. trachomatis proteins into peptide epitopes in either the cytosolic or endosomal compartments. To block presentation by the endogenous pathway, we used the trileucine peptide vinyl sulfone proteasome inhibitor, Z-L₃-VS. To inhibit the exogenous Ag-processing pathway, we used bafilomycin A1, a specific inhibitor of the vacuolar-type proton ATPase that prevents acidification of the endosomal compartment. As shown in Fig. 3⇓A, Z-L₃-VS treatment completely inhibited the ability of C. trachomatis-infected BALB/3T3 cells to stimulate the C. trachomatis-specific CD8⁺ T cell line 69. This finding was consistent with established paradigms of how CD8⁺ T cell Ags are processed by the endogenous MHC class I pathway in nonprofessional APCs (40). Treatment with the proteasome inhibitor also completely blocked C. trachomatis CD8⁺ T cell Ag presentation by infected DCs and Mφ (Fig. 3⇓, B and C). These data indicate that processing of C. trachomatis CD8⁺ T cell Ag is proteasome dependent. In contrast, treating C. trachomatis-infected DCs and Mφ with bafilomycin A1 failed to block their recognition by line 69 (Fig. 3⇓, B and C). As a positive control to test the efficacy of bafilomycin A1 in these assays, we demonstrated the ability of this drug to block anthrax toxin-mediated CD8⁺ T cell epitope delivery to the cytosol, a process that is dependent upon the acidic pH of the endosome (data not shown) (41). These results suggest that in Mφ and DCs, C. trachomatis CD8⁺ T cell epitopes are not generated inside of the endosomal compartment. However, it is also possible that the amount of C. trachomatis CD8⁺ T cell epitopes generated within this compartment is too low to stimulate IFN-γ secretion from C. trachomatis-specific CD8⁺ T cells.

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FIGURE 3.

The proteasome is required to process C. trachomatis Ag in infected APCs. BALB/3T3 (A), BALB/c DC (B), or BALB/c Mφ (C) were infected with C. trachomatis, incubated for 20 h, and then treated with DMSO alone (▪), Z-L₃-VS (▨), or bafilomycin A1 (□) for an additional 4 h. The cells were washed thoroughly and then incubated with the CD8⁺ T cell line 69 for 16 h. The amount of IFN-γ secreted into the culture supernatant was measured by ELISA. Graphs are representative of three separate assays. Error bars represent SD of triplicate wells.

Hematopoietic cells are required for priming Cap1-specific CD8⁺ T cells

Although pAPCs are believed to prime the majority of CD8⁺ T cells, nonhematopoietic cells, like fibroblasts, have been shown to prime some viral- and tumor-specific CD8⁺ T cell responses (30, 31). We found that C. trachomatis infection of fibroblasts is very productive in vitro (Fig. 1⇑A), and others have observed that epithelial cells are the major reservoir of C. trachomatis infection in vivo (18). Because high Ag level has been shown in some cases to overcome the requirement for costimulation during priming of CD8⁺ T cells (42), we hypothesized that infected nonhematopoietic cells might be sufficient for priming CD8⁺ T cells specific for the C. trachomatis CD8⁺ T cell epitope, Cap1_139–147. We used radiation bone marrow chimeras to differentiate between the ability of hematopoietic and nonhematopoietic cells to present the Cap1_139–147 epitope to naive CD8⁺ T cells. Lethally irradiated CbyB6F₁/J (CB6) mice (F₁ progeny of BALB/c and C57BL/6 mice) were reconstituted with bone marrow from BALB/c mice, C57BL/6 (B6) mice, or CB6 mice. Because the Cap1_139–147 epitope binds the H-2K^d molecule (16), the cells from mice reconstituted with BALB/c (H-2^d) bone marrow (BALB/c→CB6) can present Cap1_139–147 on both hematopoietic and nonhematopoietic cells. In contrast, the CB6 mice reconstituted with bone marrow from the C57BL/6 mice (B6→CB6) only express the H-2^d MHC allele on nonhematopoietic cells. As a result, in the B6→CB6 chimeric mice, Cap1_139–147 can only be presented on nonhematopoietic cells. Mock chimeras reconstituted with CB6 bone marrow (CB6→CB6) were also generated to control for any unexpected effects of chimerism.

All chimeric mice were infected i.p. with 1 × 10⁸ IFU of C. trachomatis. The infected mice were then allowed to recover for 2 wk before their spleens were removed and cultured with BALB/c splenocytes pulsed with Cap1_139–147 peptide. After 5 days of culture in vitro, the splenocytes from each chimera were tested in a cytotoxicity assay (Fig. 4⇓A). We found that T cells from the BALB/c→CB6 and CB6→CB6 chimeras specifically recognized P815 cells pulsed with the Cap1_139–147 peptide. This outcome was expected because both hematopoietic and nonhematopoietic cells can present the Cap1_139–147 peptide in these chimeras. However, T cells from the B6→CB6 chimera, in which only nonhematopoietic cells presented Cap1, were not able to specifically lyse Cap1_139–147 peptide-pulsed P815 cells. These findings suggest that Cap1 must be presented on hematopoietic cells to stimulate Cap1-specific CD8⁺ T cells.

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FIGURE 4.

Presentation by hematopoietic cells is essential for priming Cap1_139–147-specific CD8⁺ T cells. A, Chimeric mice were infected i.p. with 1 × 10⁸ IFU of C. trachomatis and sacrificed 2 wk after immunization. Splenocytes from infected animals were stimulated with Cap1_139–147-coated BALB/c spleen cells. Five days later, the cultures were assayed using a standard ⁵¹Cr release assay in which the target cells were P815 cells pulsed with Cap1_139–147 peptide (▪) or P815 cells alone (▨). B, Chimeric mice were infected i.p. with 1 × 10⁸ IFU of C. trachomatis, or C, injected with 5 × 10⁵ Cap1_139–147 peptide-coated BALB/c DCs, and their spleens were harvested 7 days after immunization. The number of Cap1_139–147-specific T cells was measured using an IFN-γ ELISPOT assay in which the stimulator cells were irradiated P815 cells pulsed with Cap1_139–147 peptide. Three mice were used in each experimental group. These data represent the mean values ± the SD. Each assay was performed three times with similar results.

To confirm these results, we also quantified the level of Cap1_139–147-specific T cell priming in the chimeras using IFN-γ secretion as a measure of CD8⁺ T cell effector function. The chimeras were infected i.p. with 1 × 10⁸ IFU of C. trachomatis as in the previous experiment. After 7 days, their spleens were removed, and the number of Cap1_139–147-specific CD8⁺ T cells primed following infection was quantified using an IFN-γ ELISPOT assay. Consistent with our observation from the cytotoxicity assay, Cap1_139–147-specific CD8⁺ T cells were detected in the splenocytes from the C. trachomatis-infected BALB/c→CB6 and CB6→CB6 chimeras, but not in the B6→CB6 chimeras (Fig. 4⇑B). Again, these results confirm that hematopoietic APCs are required to prime Cap1_139–147-specific CD8⁺ T cells. An alternative explanation for the inability to detect Cap1_139–147-specific CD8⁺ T cells in the B6→CB6 chimeras is that precursors for these T cells simply did not develop in these mice. To determine whether Cap1_139–147-specific T cell precursors exist and can be primed in the B6→CB6 chimeras, Cap1_139–147 peptide-pulsed BALB/c DCs were injected into these mice as a source of hematopoietic cells expressing the H-2^d molecule. Using an IFN-γ ELISPOT, we detected priming of Cap1_139–147-specific CD8⁺ T cells in both the BALB→CB6 and B6→CB6 chimeras after immunization with Cap1_139–147 peptide-coated BALB/c DCs (Fig. 4⇑C). We believe that more Cap1-specific CD8⁺ T cells were primed in the BALB→CB6 chimeras because the injected DCs can serve as both APCs and as sources of Cap1 peptide for resident H-2^d pAPCs in these mice. These results support our conclusion that presentation by hematopoietic cells is essential for Cap1-specific CD8⁺ T cell priming during C. trachomatis infection.

Adoptive transfer of C. trachomatis-infected hematopoietic cells does not result in priming of Cap1_139–147-specific CD8⁺ T cells

Because there appeared to be a requirement for hematopoietic cells in priming Cap1_139–147-specific CD8⁺ T cells in vivo, we sought to identify the hematopoietic cell type responsible for priming these T cells. Because Mφ and DCs are the typical APCs thought to prime CD8⁺ T cells, we adoptively transferred C. trachomatis-infected BALB/c Mφ or DCs into B6→CB6 chimeric mice and evaluated the generation of Cap1_139–147-specific CD8⁺ T cells using an IFN-γ ELISPOT assay. Surprisingly, neither C. trachomatis-infected DCs nor infected Mφ primed Cap1_139–147-specific CD8⁺ T cells in the B6→CB6 chimeras (Fig. 5⇓). In contrast, C. trachomatis-infected DCs and Mφ primed Cap1_139–147-specific CD8⁺ T cells in the control CB6→CB6 chimeric mice (Fig. 5⇓). This discrepancy could be explained in one of two ways. The most obvious explanation is that the C. trachomatis-infected Mφ and DCs are directly priming Cap1_139–147-specific CD8⁺ T cells in the CB6→CB6 mice. This explanation seems unlikely because the same C. trachomatis-infected Mφ and DCs failed to prime Cap1-specific T cells in the B6→CB6 chimeras. Alternatively, the infected Mφ and DCs may have lysed in vivo and released EBs that subsequently infected the CB6→CB6 mouse directly. To determine whether the priming of Cap1_139–147-specific CD8⁺ T cells following adoptive transfer of C. trachomatis-infected Mφ and DCs in the CB6→CB6 mice was the result of spread of infection from the transferred pAPCs, we repeated the experiment again while administering the antibiotic tetracycline to the recipient mice. Control mice also received C. trachomatis-infected Mφ or DCs, but were injected with PBS instead of tetracycline. We found that administration of tetracycline to CB6→CB6 mice eliminated priming of Cap1_139–147-specific CD8⁺ T cells following immunization with C. trachomatis-infected Mφ and DCs (Fig. 5⇓). Because blocking the spread of infection prevented Cap1-specific T cell priming in these mice, we conclude that adoptive transfer of C. trachomatis-infected Mφ or DCs is not sufficient for priming Cap1_139–147-specific CD8⁺ T cells.

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FIGURE 5.

Immunization with C. trachomatis-infected DC or Mφ does not prime Cap1_139–147-specific CD8⁺ T cells. BALB/c DCs (A) or BALB/c Mφ (B) were infected in vitro with C. trachomatis for 16 h, and then 5 × 10⁵ infected cells were adoptively transferred i.v. into either B6→CB6 chimeric (▪) mice or CB6→CB6 chimeric mice (▨). In one group of CB6→CB6 chimeras, mice were treated with tetracycline before and following adoptive transfer of cells (□). The spleens from the immunized mice were harvested 7 days after adoptive transfer and incubated with irradiated P815 cells pulsed with Cap1_139–147 peptide. The number of Cap1_139–147-specific CD8⁺ T cells was measured using an IFN-γ ELISPOT assay. Each group consisted of three experimental mice. Bars represent the average number of IFN-γ-secreting cells from each group ± SD.

C. trachomatis-infected hematopoietic cells are unable to stimulate naive CD8⁺ T cells in vitro

Our observation that C. trachomatis-infected Mφ did not directly prime Cap1_139–147-specific CD8⁺ T cells in vivo (Fig. 5⇑) was not surprising considering both the poor ability of C. trachomatis to replicate within these cells and their weak stimulation of C. trachomatis-specific CD8⁺ T cells in vitro (Figs. 1⇑C and 2⇑C). However, because we found that C. trachomatis-infected DCs were much better than Mφ in stimulating C. trachomatis-specific CD8⁺ T cells in vitro (Fig. 2⇑B), we anticipated that DCs might be able to prime these T cells in vivo. Our inability to detect priming of Cap1_139–147-specific CD8⁺ T cells by C. trachomatis-infected DCs could simply be attributed to our failure to transfer sufficient numbers of infected DCs to the chimeras. Because transfer of greater numbers of DCs proved lethal to the animals, we decided to address this concern by performing an in vitro priming assay in which we investigated the ability of C. trachomatis-infected DCs to stimulate proliferation of naive CD8⁺ T cells. This assay allowed us to significantly increase the E:T ratio of naive CD8⁺ T cells to C. trachomatis-infected DCs.

We measured the proliferation of CD8⁺ T cells using a CFSE dye dilution assay. Naive BALB/c splenocytes were CFSE labeled and coincubated with DCs either externally loaded with Cap1_139–147 peptide or infected with C. trachomatis. DCs that were pulsed with Cap1_139–147 peptide were also pretreated with LPS to up-regulate surface expression of costimulatory molecules. As shown in Fig. 6⇓C, DCs treated with LPS and pulsed with 10 μM Cap1_139–147 peptide stimulated proliferation of naive CD8⁺ T cells. In contrast, C. trachomatis-infected DCs failed to stimulate proliferation of naive CD8⁺ T cells (Fig. 6⇓A). As a positive control, we demonstrated that effector CD8⁺ T cells from C. trachomatis-immune mice proliferated in response to both 10 μM Cap1_139–147 peptide-coated DCs (Fig. 6⇓D) and C. trachomatis-infected DCs (Fig. 6⇓B).

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FIGURE 6.

Naive CD8⁺ T cells are not stimulated by DCs infected with C. trachomatis. Splenocytes from naive BALB/c mice (A, C, E, and G) or C. trachomatis-immune BALB/c mice (B, D, F, and H) were labeled with CFSE and cocultured with either BALB/c DCs infected with C. trachomatis (A and B) or mature DCs pulsed with 10 μM (C and D), 100 nM (E and F), or 1 nM (G and H) synthetic Cap1_139–147 peptide. After 3 days of culture, the cells were harvested, stained with anti-CD8 and anti-TCR-β Abs, and analyzed by flow cytometry. The CFSE-staining profile of the CD8⁺TCR⁺ cells is shown. The data shown are representative of three independent experiments.

The observation that C. trachomatis-infected DCs fail to prime CD8⁺ T cells in vitro is surprising and raises the possibility that there is something inherently altered in the C. trachomatis-infected DC that precludes its ability to stimulate naive CD8⁺ T cells. It is possible that C. trachomatis-infected DCs either present inadequate amounts of C. trachomatis Ag or express insufficient levels of costimulatory molecules. To address the issue of Ag presentation, we titrated the amount of peptide used to externally load LPS-matured DCs. Reducing the levels of peptide by even 10-fold eliminated the in vitro priming of naive CD8⁺ T cells (Fig. 6⇑, C, E, and G), but did not prevent the recognition of these peptide-coated DCs by effector CD8⁺ T cells from a C. trachomatis-immune spleen (Fig. 6⇑, D, F, and H). This result suggests that the level of Ag presented by C. trachomatis-infected DCs may be too low to prime naive CD8⁺ T cells but is sufficient to stimulate effector CD8⁺ T cells.

Costimulation is also important in the activation of naive CD8⁺ T cells, as determined by the inability of Cap1_139–147 peptide-coated immature DCs to prime Cap1_139–147-specific CD8⁺ T cells (data not shown). As shown in Fig. 7⇓, A–C, C. trachomatis infection of DCs results in their up-regulation of surface expression of MHC class I and II molecules as well as the accessory signaling molecule, CD40. We observed similar levels of up-regulation of these surface molecules after LPS treatment of immature DCs (Fig. 7⇓, A–C). However, there was considerably less up-regulation of surface expression of the costimulatory molecule CD86 after C. trachomatis infection compared with the up-regulation observed following LPS treatment of the DCs (Fig. 7⇓D). More dramatically, following C. trachomatis infection, DCs fail to up-regulate expression of the costimulatory molecule CD80 (Fig. 7⇓E). Taken together, these results suggest that a combination of low Ag level and impaired up-regulation of costimulatory molecule expression contributes to the inability of directly infected DCs to prime a C. trachomatis-specific CD8⁺ T cell response.

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FIGURE 7.

C. trachomatis infection results in incomplete DC maturation. After 7 days of in vitro culture, BALB/c DCs were infected with C. trachomatis (thick solid line), treated with LPS (thin solid line), or mock infected (thin dotted line). The cells were harvested 16 h after treatment. The histograms represent a gated population of DCs (CD11b⁺CD11c⁺ cells) stained with Abs specific for I-A^d (A), K^d (B), CD40 (C), CD86 (D), and CD80 (E). Ten thousand total events were counted for each sample. The data shown are representative of two independent experiments.

C. trachomatis Ag can be cross-presented by DCs in vitro

Our findings suggest that not only does C. trachomatis replicate poorly within DCs, but it also inhibits DC maturation. These observations led us to question how DCs could obtain sufficient C. trachomatis Ag to stimulate naive CD8⁺ T cells without suffering from the negative effects of infection. One mechanism by which DCs could acquire C. trachomatis Ag without direct infection is to phagocytose infected cells. After digesting C. trachomatis-infected cells, DCs could cross-present the Ag produced in those cells to CD8⁺ T cells. Cross-presentation of infected cells by DCs has been demonstrated with both viral pathogens such as vaccinia virus (43) as well as bacterial pathogens like Salmonella (44), but has not yet been shown for C. trachomatis-infected cells. To test whether DCs can cross-present Ag produced in other C. trachomatis-infected cells, we coincubated BALB/c DCs with C. trachomatis-infected MHC-mismatched (H-2^b) 1308.1 epithelial cells, C57BL/6 Mφ or C57BL/6 DCs. The antibiotic chloramphenicol was included during the coculture of donor (C. trachomatis-infected H-2^b cells) and recipient cells (H-2^d DCs) to prevent direct C. trachomatis infection of the recipient DCs. As shown in Fig. 8⇓, DCs were capable of cross-presenting Ag produced in all three types of cells infected with C. trachomatis. We observed that DCs cross-presented C. trachomatis-infected DCs (Fig. 8⇓B) more efficiently than they cross-presented infected epithelial cells (Fig. 8⇓A) or Mφ (Fig. 8⇓C). It is possible that DCs serve as the best Ag donors in this assay not because of Ag quantity, but because it is easier to engulf nonadherent DCs than to engulf adherent epithelial cells or Mφ. These data suggest that cross-presentation of C. trachomatis-infected cells is one mechanism by which DCs may stimulate C. trachomatis-specific CD8⁺ T cells.

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FIGURE 8.

DCs and Mφ are capable of cross-presenting C. trachomatis-infected cells in vitro. H-2^b APCs (1308.1 epithelial cells (A), C57BL/6 DCs (B), and C57BL/6 Mφ (C)) were infected with C. trachomatis for 24 h and then cocultured with H-2^d DCs (▨) at a ratio of 1:1 for 16 h in the presence of chloramphenicol. As a negative control, the H-2^b APCs were also cultured alone in chloramphenicol (▪). The C. trachomatis-specific CD8⁺ T cell line 69 was then added to the APCs at an E:T ratio of 10:1. After 12 h of T cell stimulation, the culture supernatant was collected and assayed for IFN-γ levels by ELISA. The data shown are representative of three independent experiments.