* Corresponding author. Mailing address: University of British Columbia Centre for Disease Control, 655 West 12th Avenue, Vancouver, BC V5Z 4R4, Canada. Phone: (604) 660-2626. Fax: (604) 660-6066. E-mail: ac.cdccb@mahnurb.trebor
Received 2007 Nov 30; Revised 2008 Jan 8; Accepted 2008 Mar 17. Copyright © 2008, American Society for MicrobiologyDendritic cells (DCs) appear to orchestrate much of the immunobiology of Chlamydia infection, but most studies of Chlamydia-DC interaction have been limited by the availability and heterogeneity of primary bone marrow-derived DCs (BMDCs). We therefore evaluated the immunobiology of Chlamydia muridarum infection in an immortal DC line termed JAWS II derived from BMDCs of a C57BL/6 p53-knockout mouse. JAWS II cells were permissive to the developmental cycle of Chlamydia. Infection-induced cell death was 50 to 80% less in JAWS II cells than in BMDCs. Chlamydia infected JAWS II cells and yielded infectious progeny 10-fold greater than that with primary BMDCs. JAWS II cells showed an expression pattern of cell activation markers and cytokine secretion following Chlamydia infection similar to that of primary BMDCs by up-regulating the expression of CD86, CD40, and major histocompatibility complex class II and secreting significant amounts of interleukin-12 (IL-12) but not IL-10. JAWS II cells pulsed with Chlamydia stimulated immune CD4 + T cells to secrete gamma interferon. Adoptive transfer of ex vivo Chlamydia-pulsed JAWS II cells conferred levels of immunity on C57BL/6 mice similar to those conferred by primary BMDCs. Taken together, the data show that JAWS II cells exhibit immunobiological characteristics and functions similar to those of primary BMDCs in terms of Chlamydia antigen presentation in vitro and antigen delivery in vivo. We conclude that the JAWS II cell line can substitute for primary BMDCs in Chlamydia immunobiological studies.
Chlamydia trachomatis is a major human pathogen which remains a grave public health threat in part because treatment-based control programs are failing to control the spread of infection (7). Chlamydia, an obligate intracellular bacterial parasite, has a unique biphasic 48- to 72-h developmental cycle with two distinct forms, an infectious but metabolically inactive elementary body (EB) and a noninfectious but metabolically active reticulate body (7). T-cell immune responses appear to be particularly important to Chlamydia immunity, and vaccination is an ideal approach to prevent infection (31). Developing an effective vaccine, however, will require identifying candidate T-cell antigens that elicit protective cellular immune responses and antigen delivery systems that evoke T-cell immunity at the tissue level.
Dendritic cells (DCs) are found in both lymphoid and nonlymphoid tissues, including mucosal surfaces, where they play an essential role as potent antigen-presenting cells (34). DCs are central to the induction of both T- and B-cell immunity in vivo (32). Antigen presentation by immature DCs induces immune tolerance (11), whereas mature DCs induce antigen-specific protective immunity through polarization of Th1 immune responses (36). Because of these unique functions, DCs have been employed in mouse model systems for vaccination and immunotherapy against cancers and infectious diseases (2), including Chlamydia infection (35). Both in vitro and in vivo studies have demonstrated the key roles played by bone marrow-derived DCs (BMDCs) in presenting Chlamydia muridarum antigens and in establishing immunity against Chlamydia infection (6, 29). Adoptive transfer with ex vivo Chlamydia antigen-pulsed BMDCs induced CD4 + Th1-cell-dominant immune responses and produced large amounts of interleukin-12 (IL-12), which stimulated CD4 + T-cell gamma interferon (IFN-γ) production, in both the murine lung model (18) and the genital model (35) of Chlamydia infection. Thus, DCs represent a powerful antigen delivery tool for the study of protective immunity against Chlamydia in vivo and provide a way to screen for and identify Chlamydia T-cell antigens in vitro, which would be useful in the design of molecular Chlamydia vaccines.
BMDCs freshly derived from mouse bone marrow are heterogeneous and are generally contaminated with macrophages and other cells. As well, BMDCs are known to be easily activated in response to in vivo conditions of the donor and ex vivo handling effects during processing, culture, and purification. Different DC generation protocols appear to produce different BMDC maturation and differentiation pathways, and therefore, the maturation condition of each BMDC generation has to be evaluated by fluorescence-activated cell sorting (FACS) analysis using activation markers. Moreover, for immunoproteomic experiments large numbers of mice have to be sacrificed to generate enough primary BMDCs in order to purify sufficient antigen-loaded major histocompatibility complexes (MHCs) for mass spectrometry. These limitations could be overcome with an immortal DC line if it should have stable characteristics and be able to grow at high density, thus satisfying quantitative requirements and providing standardized quality (8, 15, 22).
JAWS II is an immortalized immature DC line which was established from the bone marrow cultures of p53 −/− C57BL/6 mice (20). This cell line has been used for studies of antitumor (15, 24, 39) and pathogen-specific (23) immunity. In this study, we compared the characteristics of JAWS II cells to those of primary BMDCs and investigated their capacities to present Chlamydia antigens in vitro and in vivo in order to determine their potential for Chlamydia vaccine and antigen discovery studies.
Iscove's modified Dulbecco's medium (IMDM), Eagle minimum essential medium (MEM), penicillin and streptomycin, gentamicin, 2-mercaptoethanol (2-ME), sodium pyruvate, and lipopolysaccharide (LPS) were purchased from Sigma (St. Louis, MO); fetal calf serum (FCS) and l -glutamine were purchased from Gibco (Grand Island, NY); and murine granulocyte-macrophage colony-stimulating factor (GM-CSF) was purchased from R&D Systems (Minneapolis, MN). IL-4-producing hybridoma X63 was kindly provided by F. Melchers, Basilea Institute, Switzerland. All antibodies (Abs) used for FACS, enzyme-linked immunosorbent assay (ELISA), and enzyme-linked immunosorbent spot assay (ELISPOT) were obtained from Pharmingen (Mississauga, Ontario, Canada).
The JAWS II cell line, a GM-CSF-dependent DC line established from bone marrow cells of a p53-knockout C57BL/6 mouse (20), was purchased from the American Type Culture Collection (CRL-1194; ATCC, Manassas, VA). Cells were grown in a CO2 incubator at 37°C and 5% CO2 in complete culture medium consisting of IMDM with 10% FCS, 4 mM l -glutamine, 10 U/ml penicillin and 100 μg/ml streptomycin, 0.5 mM 2-ME, 1 mM sodium pyruvate, and 5 ng/ml murine GM-CSF. The medium was placed into the incubator for at least 15 min to allow it to reach its normal pH (7.0 to 7.6) before cells were added. Cultures were maintained by transferring nonadherent cells to a centrifuge tube and treating attached cells with 0.25% trypsin-0.03% EDTA (Gibco) at 37°C for 5 min, followed by pooling the two populations of cells together and dispensing them into new flasks.
The protocol for generation and purification of BMDCs used in this study has been described previously (41). Briefly, bone marrow cells were flushed from the femurs of 8- to 10-week-old female C57BL/6 mice and cultured in 150- × 15-mm Falcon petri dishes (Becton Dickinson Labware, Franklin Lakes, NJ) at 1 × 10 7 in 20 ml of IMDM supplemented with 10% FCS, 4 mM l -glutamine, 10 U/ml penicillin, 100 μg/ml streptomycin, 0.5 mM 2-ME, 10 ng/ml GM-CSF, and 5% IL-4 culture supernatants of hybridoma X63. On day 4, fresh GM-CSF (5 ng/ml) was added to the cultures. After 7 days of culture, the nonadherent cells were harvested and used as BMDCs for adoptive transfer experiments. For in vitro experiments, cells were purified using anti-CD11c magnetic beads (Miltenyi Biotec Ltd., Auburn, CA). Routinely the purity of CD11c + cells from the nonpurified population was 53.5% ± 5.2% and that of cells from the purified population was 95% ± 5% (data not shown) as determined by FACS.
Female C57BL/6 mice were purchased from Charles River (St. Constant, Canada). The mice were housed in a pathogen-free animal facility and used at 8 to 10 weeks of age. All experimental procedures were performed in accordance with the guidelines approved by the animal care committee of the University of British Columbia.
C. muridarum mouse pneumonitis (MoPn) strain Nigg was grown in HeLa 229 cells in MEM containing 10% FCS. All media for culture with live EBs used 50 μg/ml gentamicin instead of penicillin and streptomycin. EBs were purified from HeLa cells by discontinuous density gradient centrifugation as previously described (9). Purified EBs were stored in sucrose-phosphate-glutamic acid (SPG) buffer at −80°C. Inclusion-forming units (IFU) as the measure of infectivity were determined by immunostaining as previously described (40). Live EBs were heat killed (HK) by incubation in a water bath at 56°C for 30 min. To ensure that heat-treated EBs were completely inactivated, HK EBs were inoculated onto monolayers of HeLa 229 cells. No recoverable IFU were found (data not shown). The number of IFU in the original purified Chlamydia stocks was used to calculate the ratio for either live EBs or HK EBs. For all experiments unless otherwise indicated, cells were treated with either live EBs or HK EBs at a multiplicity of infection (MOI) of 3.
JAWS II cells or purified BMDCs were plated at 1 × 10 5 cells per well onto a 24-well plate containing a 12-mm coverslip and exposed to live EBs in a CO2 incubator for 24 h. After being washed with phosphate-buffered saline (PBS; Sigma), the cells were fixed in 400 μl of IC fixation buffer (eBioscience, San Diego, CA) per well at room temperature (RT) for 30 min followed by washing with PBS and rinsing with 1× permeabilization buffer (PB; eBioscience) before blocking with 1:250 naïve rabbit serum in PB (NRS/PB) at RT for 30 min. After 1 h of incubation with 1:500 mouse anti-EB serum in NRS/PB at RT, the cells were washed with PB and then stained with 1:200 Cy5-conjugated rabbit anti-mouse immunoglobulin G (IgG; Jackson ImmunoResearch, West Grove, PA) in NRS/PB at RT for 1 h, followed by washing with PB and staining with 1:200 anti-mouse CD11b-fluorescein isothiocyanate (FITC)-conjugated monoclonal Ab in NRS/PB at RT for another 1 h. Following washing with PB and rinsing with PBS, coverslips were removed and inverted onto a microscope slide in a drop of Prolong antifade kit (Invitrogen). Fluorescence was visualized by confocal microscopy (Leica Microsystems Inc., Exton, PA) using a 63× objective lens. Anti-EB sera were generated by intranasally immunizing mice twice with 1,000 IFU of live C. muridarum at 4-week intervals. Sera were collected and titrated.
Phenotypic analysis of cells was performed on a FACSCalibur sorter (Becton Dickinson, San Jose, CA) using CellQuest software (BD Bioscience, Mississauga, Ontario, Canada). Cells were incubated with medium alone, LPS (1 μg/ml), HK EBs, and live EBs for 24 h and collected. After being washed with FACS buffer (2% FCS, 0.1% NaN3 in PBS), the cells were blocked with anti-mouse CD16/CD32 (2.4G2) on ice for 15 min followed by washing and dispensed as 2 × 10 5 cells in 50 μl of FACS buffer per well into U-bottomed 96-well plates. The cells were stained on ice for 30 min using FITC-conjugated anti-mouse CD11b (M1/70), CD86 (GL1), CD80 (16-10A1), CD40 (HM40-3), MHC II (2G9), MHC I (AF6-88.5), ICAM-1 (3E2), or phycoerythrin-conjugated anti-CD11c (HL3) at 0.5 μg per well. After being washed once with FACS buffer containing propidium iodide (PI) to stain dead cells, each sample was resuspended in 0.2 ml of PBS and subjected to FACS.
The viability of cells was calculated by trypan blue exclusion, and cell death was determined by FACS using an annexin V-FITC apoptosis detection kit (Sigma) as described by the manufacturer.
Cells were plated into U-bottomed 96-well plates at 2 × 10 5 cells in 200 μl of complete IMDM per well and pretreated with medium alone, LPS, HK EBs, or live EBs for 24 h. The supernatants were discarded, and the pretreated DCs were cocultured with 5 × 10 5 Chlamydia-sensitized CD4 + T cells in 200 μl of fresh medium per well for 48 h. The culture supernatants were collected and stored at −20°C until they were tested. Chlamydia-specific CD4 + T cells were generated by intraperitoneally immunizing mice with 1 × 10 6 IFU of live EBs and boosted 2 weeks later. Mice were subsequently challenged with 1 × 10 4 IFU intranasally to ensure that they were immune. T cells were purified from murine spleens by negative selection with the MACS CD4 + T-cell isolation kit (Miltenyi Biotec Inc., Auburn, CA). As a control, CD4 + T cells from naïve mice were purified in parallel. CD4 + T cells of >90% purity were obtained as measured by FACS (data not shown). Murine sera were collected from immunized mice and kept at 4°C. Cytokines (IL-12, IL-10, and IFN-γ) in culture supernatants and MoPn-specific Abs in murine sera were determined by ELISA as previously described (40). Paired anti-mouse IL-12 (p40/p70) (C15.6 and C17.8), IL-10 (JESS-2A5 and SXC-1), and IL-6 (MP5-20F3 and MP5-32C11) monoclonal Abs for ELISA were used. To measure EB-specific Ab, anti-mouse IgG (Jackson ImmunoResearch), biotin anti-mouse IgG2a (R19-15), and biotin anti-mouse IgG1 (A85-1) were used. Briefly, plates were coated with 50 μl per well of 4-μg/ml capture Abs or 2 × 10 5 IFU per well of HK EBs in bicarbonate coating buffer at 4°C overnight. After being washed with PBS containing 0.5% Tween 20, the cells were blocked with 120 μl per well of 3% bovine serum albumin (BSA) in PBS at RT for 2 h. Fifty microliters per well of 1:250-diluted sera (for anti-EB IgG2a and IgG1) and sera serially diluted from 1:500 to 1:16,000 (for anti-EB IgG) in 0.5% BSA-PBS or the culture supernatants and recombinant cytokine standards were added to plates and held at 4°C overnight. The assay was developed at RT after washing, using 50 μl per well of 2-μg/ml biotinylated detection Abs for 1 h followed by 1:1,000 streptavidin conjugated to horseradish peroxidase in 0.5% BSA-PBS for another 1 h and substrate {0.5 mg/ml ABTS [2,2′-azino-di-(3-ethylbenzthiazolinesulfonic acid)] in 0.1 M citrate buffer, pH 4.2, containing 0.03% hydrogen peroxide (H2O2)}. The reactions were read at 405 nm with a Microplate Manager (Bio-Rad Laboratories, Hercules, CA).
The ELISPOT assay was performed as previously described (21). A MultiScreen 96-well filtration plate (Millipore Corporation, Bedford, MA) was coated with 50 μl per well of 2-μg/ml anti-mouse IFN-γ in bicarbonate coating buffer at 4°C overnight. Wells were washed and blocked with 100 μl per well complete medium at 37°C for 1 h. JAWS II cells or purified BMDCs (4 × 10 4 /well) were mixed with or without HK EBs and cocultured with Chlamydia-specific CD4 + T cells or naïve CD4 + T cells (2 × 10 5 /well) in a total volume of 200 μl per well at 37°C for 40 h. The assay was developed at RT using 50 μl per well of 2-μg/ml biotinylated detection Abs for 1 h, 1:1,000 alkaline phosphatase-conjugated streptavidin in PBS containing 0.1% gelatin and 0.5% Tween 20 for another 1 h, and 100 μl/well of phosphatase substrate containing 5-bromo-4-chloro-3-indolylphosphate and nitroblue tetrazolium (Sigma). The reactions were stopped by rinsing the reaction mixtures with water for 10 min, and spots were microscopically counted.
Adoptive transfer was performed according to a previously published protocol (29). Briefly, nonpurified BMDCs or JAWS II cells were incubated with HK EBs or live EBs in complete IMDM containing 5 ng/ml GM-CSF at 37°C, 5% CO2, for 40 h. The cells were then washed three times with PBS and resuspended in PBS at 5 × 10 6 cells/ml. Cells treated with medium alone were used as a negative control. Naïve recipient C57BL/6 mice were adoptively immunized by intravenous injection into the tail veins with 1 × 10 6 cells in 200 μl of PBS per mouse. A booster immunization was administered 2 weeks after the initial immunization. Groups of five mice were used for each adoptive transfer experiment. On day 14 after secondary inoculation, the mice were anesthetized and challenged by intranasal inoculation with 2,000 IFU of live EBs in 40 μl of PBS. The infectivity and the protective immunity were assessed by body weight changes and MoPn lung titration.
Viable MoPn titration was performed on day 10 after challenge as previously described (18). Mice were sacrificed by cervical dislocation, and lungs were aseptically removed. The tissues were homogenized with tissue grinders in 3 ml of cold SPG buffer. Tissue homogenates were centrifuged at 1,000 × g for 10 min at 4°C to remove coarse tissue debris. The clarified suspensions were serially diluted and immediately inoculated onto HeLa 299 monolayers. For the in vitro experiment, JAWS II cells or purified BMDCs were infected with live EBs and collected at various days. After being washed twice with PBS, cells were resuspended at 1 × 10 6 /ml in SPG buffer and stored at −80°C. Before testing, cells were sonicated for 5 s with an Ultrasonic Dismembranator (model 100; Fisher Scientific, Ottawa, Ontario, Canada). The confluent monolayers of HeLa 229 cells were prepared in F-bottomed 96-well plates after an overnight culture. Cell suspensions or fresh clarified tissue supernatants were serially diluted in SPG buffer and inoculated at 100 μl per well onto HeLa 229 monolayers, which were pretreated with 30 μg/ml DEAE-dextran (Pharmacia Fine Chemicals, Uppsala, Sweden) in 100 μl of Hanks balanced salt solution (Gibco) per well at RT for 15 min followed by washing with SPG buffer. After incubation at 37°C, 5% CO2, for 2 h, the supernatants were removed from HeLa cells and 100 μl of MEM with 2 μg of cycloheximide (Sigma) per well was added. The plates were placed in a cell incubator for 48 h, after which cells were fixed in 70% methanol at RT for 30 min. Immunostaining was carried out using mouse anti-EB serum as primary Ab and horseradish peroxidase-anti-mouse IgG as secondary Ab. The color was developed using 1× diaminobenzidine-metal concentrate in stable peroxide substrate buffer (Pierce, Rockford, IL), and the number of inclusions was counted under a microscope at ×200 magnification.
It has been reported elsewhere that Chlamydia can infect and undergo limited replication in human DCs (12), murine macrophages (16), and murine BMDCs (27). In the present study, we determined whether Chlamydia could also infect and replicate in JAWS II cells. To optimize the MOI of Chlamydia for JAWS II cells and primary BMDCs, cells were infected at various MOIs and viability was determined by trypan blue exclusion at several time points. The viabilities of both JAWS II cells and BMDCs were over 90% at 48 h postinfection when an MOI of 3 or less was used (Fig. (Fig.1A). 1A ). At an MOI of 30, however, viability was dramatically reduced to less than 50% in BMDCs but not in JAWS II cells. This cytotoxicity was detected as early as 24 h and increased over 72 h postinfection. These results indicate that JAWS II cells are much more tolerant to the cytotoxicity of higher MOIs of Chlamydia than are primary BMDCs.
Survival and development of Chlamydia in JAWS II cells. (A) JAWS II cells or BMDCs were infected with C. muridarum (live EBs) at an MOI of 0, 0.3, 3, or 30 for 24 h, 48 h, or 72 h. Viability was determined by trypan blue exclusion. Pooled data from three separate experiments are shown. *, P < 0.05 versus control (MOI of 0). (B) JAWS II cells or BMDCs were infected with Chlamydia at an MOI of 0, 0.3, 3, or 30 for 72 h. Cell death was determined by annexin V-PI staining using FACS. One representative experiment of three independent experiments with similar results is shown. (C) JAWS II cells or BMDCs were incubated with Chlamydia for 24 h and stained with a combination of mouse anti-EB serum and Cy5 secondary Abs followed by staining with FITC-anti-CD11b Abs. Intracellular inclusions of C. muridarum were visualized with a Leica confocal microscope using a 63× objective lens. (D) JAWS II cells or BMDCs were infected with Chlamydia for 1, 2, 3, 4, or 7 days; sonicated; and followed by titration on HeLa cells to determine the number of recoverable IFU. Pooled data from two separate experiments are shown.
Since cell death began early and was maximal for both JAWS II cells and BMDCs at 72 h with an MOI of 30, we sought to determine the type of cell death by using annexin V-FITC/PI staining. Apoptotic cell death was defined as an annexin V- and PI-positive result, and necrotic cell death was defined as an annexin V-negative and PI-positive result. By these criteria, there was little significant difference in either apoptotic or necrotic cell death between Chlamydia-infected and noninfected JAWS II cells and BMDCs at the 72-h time point when an MOI was 3 or less. At an MOI of 30, an obvious difference in cell death was observed between JAWS II cells and BMDCs, where four times as many BMDCs as JAWS II cells underwent apoptotic cell death (Fig. (Fig.1B). 1B ). These results demonstrate that apoptosis more easily occurs in primary BMDCs than in JAWS II cells at high MOIs of Chlamydia.
From the results in Fig. Fig.1, 1 , we selected an MOI of 3 as the experimental condition for evaluating the immunobiology for JAWS II cells and BMDCs following Chlamydia infection. We next investigated whether Chlamydia forms inclusions in JAWS II cells and BMDCs and noted that inclusions were clearly visible in 10% to 15% of both DC types at 24 h postinfection as detected by confocal fluorescence microscopy (Fig. (Fig.1C). 1C ). CD11b-FITC stained the cell membrane (green), and EB-Cy5 stained the inclusion (red). From the confocal images, it was clear that Chlamydia could infect and form inclusions in JAWS II cells that were phenotypically similar to those observed in BMDCs. While inclusion formation is clearly less than that for highly permissive cell lines such as HeLa, the levels are similar to those previously reported in DCs (12, 30).
To assay Chlamydia replication in JAWS II cells, the homogenates from live EB-infected JAWS II cells were propagated on HeLa 229 cells. The recoverable IFU from DCs after infection with Chlamydia for 1, 2, 3, 4, and 7 days showed similar patterns in the two DC lines, with IFU increasing up to day 4 and then decreasing at day 7. However, the recoverable IFU from 10 5 JAWS II cells were 10 times greater than those from an equivalent number of primary BMDCs. The results demonstrate that Chlamydia bacteria undergo similar developmental cycles in the two types of DCs, although they yield infectious progeny more efficiently in JAWS II cells than in BMDCs.
JAWS II cells and BMDCs were untreated or exposed to live EBs, HK EBs, or LPS for 24 h and were followed by FACS analysis for surface activation marker expression. Under resting condition JAWS II cells showed a phenotype similar to that of BMDCs as previously reported (28), except for lower MHC II expression (62% of cells stained positive with a mean fluorescence intensity of 25.5 for JAWS II cells versus 90% of cells positive with a mean fluorescence intensity of 1,413 for BMDCs). Like BMDCs, JAWS II resting cells were an immature myeloid type of DCs as determined by low expression of CD86 and CD40; moderate expression of MHC II; and high expression of CD11b, CD11c, CD80, MHC I, and ICAM-1/CD54 (Fig. (Fig.2A 2A ).
Activation marker expression by JAWS II cells and BMDCs using FACS analysis. (A) Surface marker expression on resting JAWS II cells and BMDCs. Primary BMDCs were generated from C57BL/6 mice as described in Materials and Methods. Freshly isolated BMDCs and untreated JAWS II cells were stained for surface marker expression in parallel. The percentages of positive cells (top number in each panel) and mean fluorescence intensities (bottom number in each panel) are indicated. One representative experiment from three independent experiments with similar results is shown. (B) Activation marker expression on activated JAWS II cells and BMDCs. JAWS II cells or purified BMDCs were pulsed with live EBs at an MOI of 3, LPS at 1 μg/ml, or medium (CTRL) for 24 h, and activation markers were analyzed by flow cytometry. The percentages of positive cells are shown from pooled data of three independent experiments. *, P < 0.05 versus control.
DCs mature in distinct ways in response to different microbial products, thereby instructing alternative pathways of host immunity (33). Therefore, we determined whether exposure to Chlamydia results in the same or an alternative expression pattern for maturation markers in both types of DCs. JAWS II cells and BMDCs were exposed to Chlamydia for 24 h and then subjected to FACS analysis. LPS-treated DCs were used as a positive control. In contrast to resting cells, JAWS II cells displayed enhanced expression of maturation markers CD86, CD40, and MHC II in response to Chlamydia antigens, with live and HK Chlamydia bacteria producing similar levels of marker expression (Table (Table1). 1 ). As shown in Fig. Fig.2B, 2B , the expression patterns of the markers CD86, CD40, and MHC II in JAWS II cells were similar to those of BMDCs in response to Chlamydia, suggesting that JAWS II cells regulate activation marker expression in a manner similar to that of primary BMDCs.
Surface molecular expression of activation markers by JAWS II cells a
| DC treatment | % of cells positive for surface marker (mean ± SE): | |||||
|---|---|---|---|---|---|---|
| CD11c | CD86 | CD80 | CD40 | MHC II | ICAM-1 | |
| Control | 85.1 ± 2.3 | 25.8 ± 1 | 91.1 ± 1.7 | 25.4 ± 2.2 | 56.3 ± 4.9 | 98.4 ± 0.8 |
| HK EBs | 80.4 ± 8 | 36.7 ± 0.9* | 96.9 ± 1.8 | 50.9 ± 6.5* | 71.4 ± 3.9* | 98.4 ± 0.3 |
| Live EBs | 84.9 ± 2.4 | 39.4 ± 2.2* | 94.5 ± 6.1 | 45.9 ± 3.9* | 87.1 ± 4.7* | 98.7 ± 0.8 |
| LPS | 81.7 ± 0.7 | 55.3 ± 5.3* | 98.4 ± 0.9 | 80.3 ± 1.8* | 89 ± 1.8* | 98.8 ± 4.3 |
a Percentages of JAWS II cells expressing the surface markers are given from pooled data of three independent experiments. Cells were treated as described in the legend to Fig. Fig.2. 2 . *, P < 0.05 versus control.
DCs play an essential role in secreting proinflammatory cytokines that drive the differentiation of T-cell precursors to Th1, Th2, Th17, or Treg phenotypes; the secretion of IL-12 is an especially potent inducer of Th1 immunity (19). Therefore, it is important to compare the cytokine profiles to determine whether JAWS II cells can function as well as primary BMDCs in the induction of a Th1 polarizing pattern. We tested the levels of secretion of IL-12 and IL-10 by JAWS II cells in response to Chlamydia and compared the levels with the levels of interleukins secreted by BMDCs. JAWS II cells were pulsed with either live or HK Chlamydia for 48 h, and culture media were assayed by ELISA for IL-12 and IL-10. LPS-treated cells were used as a positive control, and untreated cells were used as a negative control. Live EBs and HK EBs as well as LPS induced significantly increased levels of IL-12 but not IL-10 in both JAWS II cells and BMDCs (Fig. (Fig.3). 3 ). Similar secretion patterns in the two types of DCs suggest that JAWS II cells function as typical myeloid-type DCs capable of directing a Th1-cell-dominant response.
JAWS II cells and BMDCs pulsed ex vivo with Chlamydia secrete IL-12 but not IL-10. JAWS II cells or BMDCs were cultured alone as control (CTRL) or following pulsing with live EBs, HK EBs, or LPS. After a 48-h incubation, culture supernatants were collected and assayed by an ELISA for IL-12 and IL-10. Pooled data from three separate experiments are shown. *, P < 0.05 versus control.
To confirm that JAWS II cells function as professional antigen-presenting cells, we tested whether this DC line was capable of presenting Chlamydia antigens to Chlamydia-specific CD4 + T cells. JAWS II cells were pulsed with Chlamydia antigen for 24 h, followed by coculture with Chlamydia-specific CD4 + T cells for 48 h at a ratio of 1:5 (DC to CD4 + T cells). The coculture media were analyzed by ELISA for IFN-γ, an indicator of CD4 + Th1 immune phenotype, and for IL-10, an indicator of CD4 + Th2 immune phenotype. Cocultures of Chlamydia immune CD4 + T cells with LPS-pulsed or untreated JAWS II cells were used as negative controls. Compared with controls, HK EBs and live Chlamydia-pulsed DCs from either source induced immune splenocytes to produce significantly higher levels of IFN-γ but not IL-10 (Fig. (Fig.4A). 4A ). LPS-treated DCs produced large amounts of IL-12 (Fig. (Fig.3). 3 ). However, immune CD4 + T cells failed to produce IFN-γ when cocultured with LPS-treated DCs, confirming that the high secretion of IFN-γ was due to antigen-specific recognition.
Chlamydia-pulsed JAWS II cells and BMDCs present antigens to Chlamydia-specific CD4 + T cells. (A) JAWS II cells and BMDCs were pulsed with medium as control (CTRL), live EBs, HK EBs, or LPS for 24 h. Splenic CD4 + T cells were isolated from Chlamydia-infected mice or naïve control mice and cocultured with pretreated JAWS II cells or BMDCs for 48 h. Cytokines secreted by CD4 + T cells were assayed by ELISA. Pooled data from three separate experiments are shown. *, P < 0.05 versus control. (B) JAWS II cells or BMDCs were cocultured with splenic CD4 + T cells isolated from Chlamydia-immunized mice (Imm.) or naïve control mice (Naïve), with or without HK EBs, for 24 h for ELISPOT assay. Spots were microscopically counted as IFN-γ-secreting cells. Pooled data from two separate experiments are shown. *, P < 0.05 versus control (HK EB − ).
An ELISPOT assay was next carried out to quantitate the antigen presentation function of JAWS II cells. JAWS II cells were mixed with or without HK EBs and cocultured with Chlamydia-specific CD4 + T cells for 24 h at the same ratio that was used for ELISA. IFN-γ-secreting cells were counted microscopically. The coculture of sensitized CD4 + T cells with JAWS II cells or BMDCs and HK EBs produced significantly increased numbers of IFN-γ-secreting cells, compared with nonpulsed DCs or naïve CD4 + T cells (Fig. (Fig.4B). 4B ). Thus, the ELISPOT assay confirmed Chlamydia-specific IFN-γ production and revealed that JAWS II cells can function as well as primary BMDCs in antigen presentation. Collectively, these results show that JAWS II cells present Chlamydia antigens in vitro as well as primary BMDCs do and suggest that Chlamydia-pulsed JAWS II cells may induce a protective CD4 + Th1 immune response in vivo following adoptive transfer.
It has been demonstrated that adoptive immunization with ex vivo Chlamydia-pulsed BMDCs induces protective antigen-specific immune responses against Chlamydia in a murine model of infection (35). Thus, we tested whether JAWS II cells could also be used for this purpose and compared results with those for primary BMDCs. JAWS II cells were pulsed ex vivo with live and HK Chlamydia. After incubation for 40 h, Chlamydia antigen-pulsed JAWS II cells were intravenously transfused into mice. A booster immunization was given 2 weeks later. Sera were collected before challenge. To compare the protective immune responses induced by adoptive transfer using antigen-pulsed JAWS II cells, the same experiments were carried out using primary BMDCs in parallel.
Anti-Chlamydia Ab was followed as an index of immune responses following DC immunization and was titrated by ELISA using whole EB as antigen. The reaction was stopped when the positive-control optical density was at 0.5, and the titer was scored as positive if the optical density value was ≥0.2 units. The titers of anti-Chlamydia Abs from murine sera adoptively transferred with either HK- or live EB-pulsed DCs were 1:8,000 and 1:16,000, respectively (Fig. (Fig.5A). 5A ). Compared with control, specific anti-Chlamydia Abs were generated after adoptive transfer with HK EB- or live EB-pulsed DCs. These data demonstrate that mice generated Chlamydia-specific immune responses following immunization with either ex vivo Chlamydia antigen-pulsed JAWS II cells or primary BMDCs.
Protective immunity was generated following adoptive immunization with ex vivo Chlamydia-pulsed JAWS II cells and BMDCs. Groups of five mice were intravenously immunized with ex vivo JAWS II cells alone, live EBs, or HK EB-pulsed JAWS II cells. Identical immunization with BMDCs was carried out in parallel. Two weeks after initial adoptive immunization, mice were boosted by secondary immunization. (A) Sera were collected from mouse tail veins 2 weeks later and before Chlamydia challenge. An ELISA was used to titrate the levels of Chlamydia-specific IgG Ab in serum. (B) The levels of Chlamydia-specific IgG1 and IgG2a Abs in serum were measured by ELISA, and mean optical density at 415 nm was used to calculate the ratio of IgG2a to IgG1. A 1:250 dilution of serum was used for all assays. (C) Mouse body weight changes after Chlamydia challenge infection. Two weeks after the second immunization, mice were intranasally challenged with 2,000 IFU of live Chlamydia bacteria, and body weight was monitored on selected days. (D) Ten days after Chlamydia challenge, the lungs were isolated and Chlamydia IFU were quantitated on HeLa cells. A representative experiment from two independent experiments is shown. *, P < 0.05 versus JAWS II-alone or BMDC-alone control.
We next tested IgG isotype Abs as a surrogate measure of Th polarization. The ratio of IgG2a to IgG1 Abs significantly increased after immunization with ex vivo live EB-pulsed DCs of either derivation (Fig. (Fig.5B). 5B ). These results indicate that ex vivo live EB- or HK EB-pulsed JAWS II cells induce Th1 immune responses as well as primary BMDCs do.
To validate whether the immune response induced by adoptive transfer with antigen-pulsed JAWS II cells was protective, vaccinated mice were intranasally challenged with 2,000 IFU of Chlamydia 2 weeks after the second adoptive immunization and protection was measured by monitoring weight loss and lung bacterial infectious titers. Mice receiving either JAWS II cells or BMDCs pulsed with live or HK Chlamydia showed significantly less body weight loss after challenge than did control mice (Fig. (Fig.5C). 5C ). Mice adoptively immunized with JAWS II cells or BMDCs pulsed with either live or HK Chlamydia showed significantly reduced numbers of IFU from lung homogenates (Fig. (Fig.5D). 5D ). Immunization with live EB-pulsed JAWS II cells showed nearly complete protection, whereas immunization with HK EB-pulsed JAWS II cells exhibited partial protection when mice were challenged with 2,000 live Chlamydia bacteria. Similar results were achieved with BMDCs. These results demonstrate that JAWS II cells can be used for adoptive immunization experiments and induce levels of protection equivalent to those induced by primary BMDCs.
DCs are at the heart of adaptive immune responses and are key determinants to understanding Chlamydia immunobiology (17). They are likely the cellular basis for the entanglement of Chlamydia immunity and disease pathogenesis that is so characteristic of this pathogen (28). By linking innate to adaptive immune responses, DCs regulate the antigen-specific developmental pathway of effector and memory T lymphocytes (3, 33). DCs are specialized to capture and process antigens, converting proteins to peptides that are presented on MHC molecules for recognition by antigen-specific T cells (14, 37). Fresh DCs generated from murine bone marrow have been critical to developing in vitro cell systems to study Chlamydia immunobiology (7). Important findings emerging from these studies include the role of DCs in antigen delivery for induction of protective immunity (17, 35), differential maturation of DCs in response to live versus dead organisms (28), and the ability of Chlamydia to establish chronic infection in DCs (27). However, the needed large quantities of BMDCs require sacrificing a large number of mice and performing complicated cell culture procedures. Moreover, BMDCs exhibit variable heterogeneity in developmental response, which may make results difficult to interpret and replicate. Thus, there is a need for a well-characterized BMDC line that can serve as a substitute for primary BMDCs to advance the studies of Chlamydia immunobiology. Here, we characterized an immature DC line, JAWS II, which originated from BMDCs of p53 −/− C57BL/6 mice. JAWS II cells have already been used for study of other infectious pathogens (23, 38) and compared with BMDCs (15, 22). However, they have not yet been studied for their interaction with Chlamydia.
Our previous studies have shown that Chlamydia bacteria survive within BMDCs and develop inclusions, which are able to infect HeLa cells (27). In this study, we confirmed that Chlamydia bacteria also infect JAWS II cells and exhibit a productive developmental cycle. FACS analysis supported the idea that resting JAWS II cells are immature myeloid DCs with characteristics similar to those of fresh resting BMDCs. After Chlamydia stimulation, JAWS II cells expressed maturation of activation markers and immune responses similar to those of BMDCs. Although the mean intensity of MHC II expression is much lower on resting JAWS II cells under resting conditions than on primary BMDCs, MHC II expression on both DC lines increased after either EB or LPS treatment (13.3% for primary BMDCs and 37.7% for JAWS II cells [data not shown]). Furthermore, the data for in vivo protection indicated that the lower surface expression of MHC II molecules on resting cells did not affect the function of JAWS II cells in antigen presentation.
Adoptive immunization studies revealed that JAWS II cells function as well as BMDCs do in in vivo protective immunization assays. We conclude that JAWS II cells display immunobiological characteristics and functions similar to those of BMDCs in response to Chlamydia.
JAWS II cells showed lower background levels of cytokine IL-12 and IL-10 production and in antigen presentation assays following Chlamydia stimulation. They also showed lower background and better consistency in the induction of immune responses. These observations likely reflect the fact that JAWS II cells are more homogeneous in phenotype than BMDCs. Thus, JAWS II cells may actually be a better choice than BMDCs for in vitro immunogenicity and antigen discovery and in vivo immunization experiments.
Of interest, JAWS II cells also showed less cytotoxicity than BMDCs did after high-MOI infection with Chlamydia. Interestingly, the cytotoxicity was mainly triggered via apoptosis as assayed by annexin V staining. The reduction of Chlamydia-induced cytotoxicity in JAWS II cells suggests that p53 may be involved in the undefined biochemical pathway that underlies high-MOI Chlamydia-induced cytotoxicity. p53 is a transcription factor that can induce growth arrest, apoptosis, and cell senescence (4). Upon DNA damage or other stress, p53 is activated and either induces cell cycle arrest, allowing repair of DNA and survival of the cell, or if genome damage is unrepairable, triggers apoptosis (30). Cell survival through the suppression of apoptotic cell death has been demonstrated in p53-deficient cells or p53 down-regulated cells in other systems (1, 10, 26). At low MOI it is known that Chlamydia-infected cells remain mitotically active and resist induction of apoptosis through the action of chlamydial protease/proteasome-like activity factor (CPAF) (13, 25). At a high MOI, cells undergo early cytotoxicity that has been hypothesized to be due to the Chlamydia cytotoxin (5). Our findings suggest that p53 may be involved in this response, at least in DCs.
In summary, Chlamydia can infect and productively develop in JAWS II cells. Resting JAWS II cells exhibit a resting phenotype of immature myeloid-type DCs that can be activated after interaction with Chlamydia. Chlamydia-pulsed JAWS II cells secrete increased amounts of IL-12 but not IL-10 and present Chlamydia antigen to infection-sensitized CD4 + T cells. Adoptive transfer with ex vivo Chlamydia antigen-pulsed JAWS II cells induces protective immunity. Because of these cellular characteristics and functions, JAWS II DCs are a convenient substitute for primary BMDCs in cellular and molecular studies of Chlamydia-DC immunobiology.
This work was supported by research funds from the Canadian Institute for Health Research grant 20R91961.
