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The Journal of Immunology, 2007, 179, 2713 -2721
Copyright © 2007 by The American Association of Immunologists, Inc.
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Antigen Transmission by Replicating Antigen-Bearing Dendritic Cells1

Jun Diao*,{dagger}, Erin Winter*,{dagger}, Wenhao Chen*, Feng Xu* and Mark S. Cattral2,*,{dagger}

* Toronto General Hospital Research Institute, University Health Network, and {dagger} Department of Surgery, University of Toronto, Ontario, Canada


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
During steady-state conditions, conventional spleen dendritic cells (DC) turn over every 2–3 days. Recent evidence indicates that in situ proliferation of DC arising from immediate conventional DC precursors is an important contributor to their homeostasis. In this study, we report that replication-competent conventional DC precursors and DC can internalize and transfer model particulate and soluble Ags directly to their DC progeny during cell division. Real-time confocal microscopy and flow cytometry indicated that Ag transmission to progeny was symmetrical, and suggested that other mechanisms of inter-DC Ag transfer were not involved. Soluble protein Ags inherited by DC progeny were presented effectively to Ag-specific T lymphocytes. Furthermore, we show that the number of DC, and the proportion that are actively proliferating, expands several-fold during an immune response against a viral infection. Our results point to an unanticipated mechanism in which DC are continuously replaced from Ag-bearing replication-competent precursor cells that pass Ag molecules onto their progeny through successive cell divisions. Our findings help explain how Ag may persist in a population of DC despite the brief lifespan of individual mature DC.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Dendritic cells (DC)3 are specialized APCs that have a key role in initiating and regulating adaptive immune responses and maintaining self-tolerance (1). The estimated lifespan of conventional DC in spleen during steady-state conditions is <2–3 days (2, 3, 4). The traditional view of the DC lifecycle holds that circulating precursors migrate to lymphoid and parenchymal tissues and differentiate into nonreplicating immature DC (1, 5, 6). Recent evidence indicates, however, that local proliferation of spleen DC contributes substantially to their homeostasis (7). We and others have shown that the majority of these dividing spleen DC arise from a distinct population of immediate conventional DC precursors (cDCp) (8, 9). cDCp are CD11c+, MHC class II–, lineage (CD4, CD8, B220, DX5, F4/80, CD19, CD3)–, and generate exclusively CD11c+ MHC class II+ DC that can continue to replicate for several generations.

Classic models of DC biology are based on the premise that DC have little or no capacity for cell division (1, 10). Accordingly, the entire process of Ag internalization, processing, and presentation is considered a function of single DC. The potency of DC in stimulating immune responses has been attributed, at least partly, to their inability to degrade internalized Ags rapidly, which increases the duration of Ag presentation (11, 12). In light of their apparent brief lifespan, however, this explanation for Ag persistence seems incomplete. In this study, we investigate the hypothesis that DC replication provides a novel mechanism for Ag transmission to their DC progeny, and in this way, sustains a population of DC capable of presenting Ag to circulating lymphocytes. Our findings reveal that replication-competent Ag-bearing cDCp and DC pass Ag molecules onto their progeny directly through successive cell divisions. Furthermore, we show that this mechanism of Ag transmission is operational during an inflammatory response induced by adenoviral infection, contributing to increased numbers of DC.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

Male C57BL/6 mice were purchased from Charles River Laboratories. C57BL/6.SJL congenic, C57BL/6-transgenic (ACTbEGFP)OPsb/J, OT-II and OT-1 trangenic mice were purchased originally from Taconic Farms or The Jackson Laboratory, and bred in our animal facility. Mice were maintained in pathogen-free conditions in accordance with institutional guidelines, and used at 6–8 wk of age.

Antibodies

Anti-CD11c (clone HL3), I-Ab (KH74, 25-9-17), CD3 (17A2), CD4 (CT-CD4), CD8{alpha} (53-6.7), CD19 (1D3), pan-NK (DX5), GR-1 (RB6-8C5), CD11b (M1/70), B220 (RA3 6B2), CD45 (30-F11), CD45.2 (104), CD45.1 (A20), CD40 (3/23), CD80 (16-10A1), CD86 (GL1), CD44 (IM7), and CD16/32 (2.4G2) were purchased from BD Pharmingen. Anti-F4/80 (A3-1) was purchased from Serotec. These Abs were either unlabeled or conjugated to FITC, PE, allophycocyanin, or biotin, as indicated. Unlabeled Abs were revealed with PE-conjugated goat anti-mouse Ig and biotinylated Abs with allophycocyanin, PC5, or PC7.

Cell isolation

DC and cDCp were isolated from bone marrow and spleen, as described (8). Briefly, spleens of 10–15 mice were minced, digested with collagenase and DNase for 0.5 h at 37°C, and incubated with EDTA. Mononuclear cells were isolated from spleen and bone marrow by Lympholyte-M (Cedarlane Laboratories) density-gradient centrifugation (13), and enriched for CD11c+ cells by positive selection using MACS (Miltenyi Biotec) and CD11c+-immunomagnetic beads. Cells retained in the column were eluted and labeled with anti-I-Ab FITC, anti-CD11c PE, and anti-lineage markers (anti-CD3, anti-CD4, anti-CD8{alpha}, anti-CD19, anti-B220, anti-F4/80 and anti-pan NK/biotin-allophycocyanin or PC7) mAbs. Lineage-negative CD11c+ MHC II– cells (i.e., cDCp) and lineage-negative CD11c+ MHC II+ DC fractions were sorted on a MoFlo High-Speed Cell Sorter using Summit acquisition and analysis software (DakoCytomation). The purity of the cell populations used was routinely ≥99% based on reanalyzed samples. OT-1 and OT-II T cells were isolated from spleen and lymph nodes by negative selection (Miltenyi Biotec).

BrdU labeling of phagocytic DC

CD45.2 mice were injected with 50 µl of 2.7% YG-latex beads in 0.3 ml of saline (0.5 µm; Polysciences). Eight hours later, BrdU (1 mg; Sigma-Aldrich) was administered i.p. every 6 h until cell recovery to ensure that it was continuously available to dividing cells (14). Purified CD45.1 DC (1 x 106 per mouse) were injected i.v. into these mice at the start of BrdU treatment. Bone marrow and spleen were recovered for analysis of bead uptake and BrdU incorporation by flow cytometry, as described (8). The proportion of beads captured during the 8 h before BrdU injection was calculated by subtracting the percentage of the adoptively transferred CD45.1 DC that contained beads from the total percentage of spleen DC that contained beads at 32 h. In some experiments, bead+ DC and bead– DC were sorted for analysis of BrdU by immunohistochemistry (BD Biosciences) and confocal microscopy (Olympus FluoView 1000 LSCM).

Stromal coculture

Sorted cDCp (5 x 104) were pulsed with OVA (200–500 µg/ml; Sigma-Aldrich) or dextran conjugated with Alexa 647 or Texas Red (1 mg/ml; m.w., 10,000; Molecular Probes) for 1–2 h, labeled with CFSE (Molecular Probes), and cultured on a confluent monolayer of irradiated (25 Gy) stromal cells derived from S17 cells (a gift from K. Dorshkind, University of California, Los Angeles, CA (15)). The cells were cultured in RPMI 1640 supplemented with 10% FBS, 50 µM 2-ME, 1 mM sodium pyruvate, 10 mM nonessential amino acids, 50 U/ml penicillin, and 50 µg/ml streptomycin (complete medium) in the presence of GM-CSF (1000 U/ml; BD Pharmingen), as described (8). At 3 days, the monolayer was disrupted with 0.25% trypsin/1 mM EDTA and repeated pipetting. The recovered cells were washed and stained with mAbs for flow cytometry or cell sorting.

Real-time confocal microscopy

cDCp from GFP-transgenic mice were pulsed with dextran conjugated with Texas Red and cocultured on S17 with complete medium in LabTek chamber slides. Live cell microscopy was performed using a multichannel confocal laser-scanning microscope (Olympus FluoView 1000 LSCM; PlanApo x60 oil-immersion objective; NA 1.4) and accompanying imaging analysis software (FV10-ASW 1.5; Olympus). The microscope is housed in chamber maintained at 37°C in an atmosphere of 5% CO2, allowing cells to remain viable for several days. Images were acquired every 10 s over several hours. The peak excitation/emission wavelengths for GFP and Texas Red were 488/526 and 516/615 nm, respectively.

Cell cycle analysis

FACS-purified OVA– DC and OVA+ DC pooled from three mice were fixed in 75% ethanol at 4°C for 16 h, and stained with propidium iodide (50 µg/ml) in PBS containing 0.1% Triton X-100 and 0.2 mg/ml RNase for 30 min at room temperature. DNA content was determined by flow cytometry using the doublet discrimination unit and analyzed by ModFit LT software (Verity Software).

Ag presentation assays

DC arising from OVA-pulsed cDCp were purified by FACS and irradiated. Graded numbers of irradiated cells were incubated with CD8+ OT-1 T cells (purity ≥95%, 5 x 104/well) for 3 days in a humidified atmosphere of 5% CO2 in air at 37°C. Proliferation was assessed by [3H]thymidine incorporation, as described (16).

Adoptive transfer studies

OVA-pulsed and CFSE-labeled cDCp (0.5–1 x 106) were injected i.v. into mice that had received 1–2 x 106 CFSE-labeled OT-1 CD8+ T cells 24 h earlier. CFSE-labeled syngenic B cells were cotransferred with cDCp in some experiments, which served as a control for nondividing cells. Spleens and lymph nodes were collected 3 days later, and DC and T cell proliferation in vivo was assessed by CFSE dilution, as described (8).

To assess the stimulatory capacity of the in vivo DC progeny derived from OVA-pulsed cDCp, 2 x 106 cDCp were injected directly into the spleen, and 3 days later, undivided CD45.2 DC, divided CD45.2 DC, and endogenous CD45.1 DC were sorted by FACS. The recovered cells were irradiated, and 2.5 x 103 were incubated with 1 x 105 OT-1 CD8+ T cells in triplicate for 3 days. T cell proliferation was evaluated by [3H]thymidine incorporation.

Adenovirus infection

Adenovirus encoding bacterial β-galactosidase (Adv-βgal) was injected i.v. into B6 mice (1 x 1010 virus particles per mouse). Preparation, purification, and titration of Adv-βgal have been described previously (17). Latex beads were injected at the same time or on the first, third, or sixth day of infection. BrdU labeling was initiated 8 h after bead injection and continued over 12 h. Spleen mononuclear cells were isolated, and the total number of DC and the proportion that were bead+ and BrdU+ were determined by flow cytometry.

Statistics

Continuous variables are expressed as mean ± SD and analyzed by two-tail Student’s t test. A p value <0.05 is considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Ag-bearing DC replicate in vivo

We first injected i.v. the commonly studied model Ag OVA or PBS control into B6 mice and treated them with BrdU for 48 h. The frequency of spleen DC that incorporated BrdU was 46 ± 6% and 31 ± 4% (p < 0.05) in OVA-treated and control mice, respectively (Fig. 1a). This finding supported the hypothesis that Ag-bearing DC or their immediate precursors could be dividing in vivo. Because BrdU uptake alone cannot establish the identity of a dividing cell population, we analyzed the cell cycle status of OVA-bearing CD11c+ lineage– spleen cells, which are comprised of DC and cDCp, 2 h after i.v. injection of OVA-conjugated with FITC or PBS control. OVA-FITC was internalized by 30% of spleen DC, consistent with previous studies (2), and by similar percentage of MHC class II– cDCp (Fig. 1b). OVA-FITC+ and OVA-FITC– spleen DC and cDCp were sorted by FACS, fixed, and stained with propidium iodide. We found that 3.7 ± 1.5% (n = 4) of OVA-FITC+ DC/cDCp and 4.0 ± 0.7% of OVA-FITC– DC/cDCp were in the S/G2/M phases of the cell cycle (Fig. 1b). These rates were similar to those in control mice injected with PBS. To verify that these OVA-FITC+ cells had the capacity to replicate, we injected sorted OVA-FITC+ DC and cDCp from CD45.1 mice into CD45.2 congenic mice and treated them with BrdU. Naive OT-1 T cells or OT-II T cells were injected i.v. into the recipient mice 24 h before DC/cDCp injection. We found that 15% of the transferred CD45.1 DC/cDCp incorporated BrdU at 24 h (Fig. 1c). At 3 days, most of the transferred OT-1 and OT-II T cells had divided, confirming that OVA-FITC+ DC/cDCp carried Ag.


Figure 1
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  		FIGURE 1. Ag-bearing DC replicate in vivo. a, Mice were immunized with OVA or PBS (control) and treated with BrdU for 48 h before cell recovery. CD11c+ lineage– spleen DC were isolated by FACS and analyzed by flow cytometry. Dot plots indicate percentage of cells expressing BrdU and MHC class II. b, Mice were injected with OVA conjugated with FITC or PBS (control). Two hours later, OVA-FITC+ and OVA-FITC– spleen CD11c+ lineage– cells were sorted using the indicated gates and stained with propidium iodide; the percentage of cells in the S/G2/M phases of the cell cycle was determined by flow cytometry and is indicated in the histograms. c, Sorted OVA-FITC+ spleen CD11c+ lineage– cells from CD45.1 mice were transferred into CD45.2 mice that had received either CFSE-labeled OT-I or OT-II T cells 24 h earlier. Recipient mice were treated with BrdU. Dot plots indicate percentage of CD45.1 DC that incorporated BrdU (left) at 24 h, and CFSE dilution of OT-1 and OT-II at 3 days. Results are representative of at least three independent experiments.

 
Labeling of replicating DC with latex beads in vivo

To further confirm that Ag-bearing DC are dividing in vivo, mice were injected i.v. with YG-labeled latex beads, which served as a model particulate Ag, and treated with BrdU (Fig. 2a). The proportion of spleen DC and cDCp that contained beads 2 h after injection was 26–32% (Fig. 2b). cDCp accounted for 18% of the total number of bead-positive CD11c+ lineage– cells in spleen. BrdU was initiated 8 h after bead injection to allow for clearance of beads from the circulation (18), and continued for 24 h before analysis. We adoptively transferred DC from congenic CD45.1 mice into each CD45.2 recipient at the start of BrdU treatment, which provided a surrogate measure of bead reuptake during the 24-h BrdU-labeling period. Previous studies established that adoptively transferred DC have the same capacity to divide and phagocytose beads and dextran as compared with endogenous DC (data not shown).


Figure 2
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  		FIGURE 2. Labeling of replicating DC with latex bead in vivo. a, Experimental protocol. CD45.2 congenic mice were injected with YG-latex beads i.v., and 8 h later were treated with BrdU for 24 h. CD45.1 congenic DC were injected i.v. into each recipient at the start of BrdU treatment. b, Detection of YG-latex beads in spleen cDCp and CD11c+ MHC class II+ DC 2 h after injection of YG-labeled beads. Numbers in histograms indicate percentage of cells. c, Analysis of bead uptake in adoptively transferred CD45.1 DC. d, Analysis of BrdU incorporation and MHC class II expression by bead+ CD11c+ lineage– spleen cells. Data from control mice that received unlabeled beads and no BrdU treatment are shown in the left dot plots. e, Proportion of bead+ and bead– CD11c+ lineage– cells in spleen and that incorporated BrdU. Error bars are mean ± SD of four independent experiments with three mice per experiment. f, Immunofluorescent microscopy of FACS-purified spleen DC showing beads inside DC. The nucleus of BrdU+ DC is stained brown. Original magnification, x600. g, Confocal microscopy of a bead+/BrdU+ DC.

 
Analysis of the adoptively transferred CD45.1 DC revealed that a small percentage (3–4%) were bead+ with most containing a single bead; this finding confirmed that the vast majority of beads were rapidly cleared from circulation (Fig. 2c). The proportion of endogenous spleen DC that contained beads 32 h after bead injection was 28.1 ± 3.7% (n = 4; Fig. 2d). BrdU was incorporated by 15 ± 4% and 21 ± 3% of bead+ and bead– spleen DC, respectively (Fig. 2e). Fluorescent and confocal microscopy of sorted DC verified that the beads were inside BrdU+ DC and not simply attached to their cell surface (Fig. 2, f and g). Collectively, these findings indicate that internalized beads captured by replicating DC can be transmitted to their daughters.

Replicating Ag-bearing DC arise from cDCp

Our finding that cDCp could internalize i.v. injected OVA and latex beads suggested that they were the source of dividing Ag-bearing DC in vivo. To test this possibility, sorted bone marrow or spleen cDCp and DC from CD45.2 mice were labeled with CFSE, pulsed with OVA, and injected i.v. into nonirradiated CD45.1 mice that had received CFSE-labeled OT-I T cells 24 h earlier. At 3 days, all of the cells arising from bone marrow cDCp expressed MHC class II, and 50–60% had divided, as determined by CFSE dilution and BrdU uptake (Fig. 3a). Similar results were obtained with sorted spleen cDCp. Cotransferred CFSE-labeled B cells provided a negative control for cell division. As anticipated, transferred mature DC were not detectable in spleen beyond 48 h (data not shown). OT-1 T cells had completed several rounds of division at 3 days, indicating that the DC arising from cDCp could present OVA (Fig. 3b). Cross-presentation of OVA by endogenous DC was not involved because OT-1 T cell proliferation was not stimulated by OVA-pulsed cDCp from Bm1 mice (data not shown).


Figure 3
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  		FIGURE 3. Progeny of Ag-bearing cDCp can stimulate T cell proliferation. OVA-pulsed and CFSE-labeled bone marrow cDCp from CD45.2 mice were injected into CD45.1 mice that had received naive CFSE-labeled OT-1 T cells 24 h earlier. a, Flow cytometric analysis of CFSE staining and MHC class II expression in CD45.2 DC at 3 days. b, CFSE dilution and BrdU incorporation of cotransferred OVA-pulsed cDCp (left) and syngeneic B cells (right). c, CFSE dilution of OT-1 T cells at 3 days in mice injected with OVA-pulsed (left) or BSA-pulsed cDCp (right) at 3 days. d, The undivided and divided DC progeny of OVA-pulsed and CFSE-labeled CD45.2 cDCp and CD45.1 endogenous DC were FACS purified using the gates indicated 3 days after injection, irradiated, and mixed with naive OT-1 T cells. Proliferation was measured by thymidine uptake. Results are representative of three independent experiments. Error bars are mean ± SD.

 
DC progeny of Ag-bearing cDCp stimulate T cell proliferation

Although our findings indicate that Ag-bearing cDCp generate dividing DC, it remained unclear whether their progeny could stimulate T cells. We therefore sorted divided and undivided CD45.2 DC based on CFSE expression 3 days after adoptive transfer. The cells were irradiated, and 2.5 x 103 were cultured with 1 x 105 OT-1 CD8+ T cells. We found that both populations were capable of cross-presenting OVA peptide (Fig. 3c). As expected, DC recovered from mice that received BSA-pulsed CFSE-labeled CD45.2 cDCp and endogenous CD45.1 DC did not stimulate OT.1 CD8+ T cells.

Replicating DC transmit internalized Ag directly to their progeny

DC acquire Ags primarily by endocytosis and phagocytosis (19, 20). Ags can also be transferred between DC through various mechanisms, including uptake of vesicular exosomes and apoptotic bodies, exchange of cell membrane, and transfer of organelles and peptides via tunneling nanotubules and gap junctions (21, 22, 23, 24, 25). To further define how daughter DC acquire Ag, we used an in vitro coculture system developed in our laboratory that promotes survival and proliferation of DC from cDCp (8). cDCp from CD45.2 mice were pulsed with dextran conjugated with a fluorophore (Alexa 647 or Texas Red), labeled with CFSE to monitor cell division, and cultured on S17 stromal cells in the presence of GM-CSF for 3 days (Fig. 4a). Dextran is a model polysaccharide Ag that is endocytosed efficiently by cDCp and DC (26). To assess dextran acquisition from the culture medium and from other inter-DC transfer mechanisms, we mixed cDCp from CD45.1 mice with the dextran-pulsed CD45.2 cDCp at a 1:1 ratio before placing them on the S17 monolayer. Confocal microscopy confirmed that CD45.1 DC were contiguous with CFSE-stained CD45.2 DC on the monolayer (data not shown). Flow cytometry and fluorescent microscopy of recovered CD45.2 DC showed that internalized dextran was passed on to DC progeny. The amount of intracellular dextran decreased by about half with each division cycle, as determined by mean fluorescence intensity (Fig. 4, b and c). By contrast, the vast majority of control CD45.1 cDCp and their progeny neither contained dextran nor acquired the CFSE stain. These results indicate that dextran internalized by proliferating DC is transmitted efficiently and directly to their progeny.


Figure 4
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  		FIGURE 4. Internalized Ag is transmitted directly to the progeny of replicating DC. a, Experimental protocol. CD45.2 cDCp were pulsed with dextran conjugated with a fluorophore or OVA, labeled with CFSE, mixed with control CD45.1 cDCp, and cocultured on S17 for 3 days. b, Flow cytometric analysis of dextran-Alexa 647 expression by DC progeny of CD45.2 cDCp (dark line) in each division cycle as defined by CFSE staining intensity; the amount of intracellular dextran was quantified by flow cytometry according to mean fluorescence intensity, as indicated in the histograms. The mean fluorescence intensity of CD45.1 cells (light line) in each histogram was ≤0.8. c, Confocal microscopy of FACS-purified undivided and divided DC from CD45.2 cDCp pulsed with dextran-Texas Red and control CD45.1 DC. Original magnification, x600. d, Divided and undivided progeny of OVA-pulsed CD45.2 cDCp and CD45.1 DC were purified by FACS using the indicated gates, irradiated (25 Gy), and mixed with naive OT-1 T cells for 3 days. DC pulsed with BSA served as a negative control. Cell proliferation was measured by thymidine uptake. Results are representative of four independent experiments. Error bars are mean ± SD.

 
We next sorted undivided and divided progeny of OVA-pulsed CD45.2 cDCp and control CD45.1cDCp after they were cocultured together for 3 days, and incubated them with naive CD8+ T cells from OT-I transgenic mice. Gates were adjusted on the FACS to ensure that the overlap between undivided and divided cells was <2%, and was confirmed by postsort flow cytometric analysis. The progeny of CD45.2 cDCp stimulated OT-1 T cell proliferation, although the magnitude of the response was slightly less than with equivalent numbers of undivided CD45.2 DC (Fig. 4d). By contrast, control CD45.1 DC were poor stimulators of OT-1 T cells. This finding suggests that little or no free OVA is released into the medium and that acquisition of OVA from apoptotic CD45.2 DC or other mechanisms of intercellular Ag and peptide transfer are not significant factors (22, 23, 25, 27, 28). Thus, dividing DC transmit internalized Ag directly to their progeny, which in turn can present inherited Ag to Ag-specific T cells.

Real-time confocal microscopy of DC division

To further investigate how internalized Ags are transferred to progeny, we used time-lapse confocal microscopy to visualize division of DC in real time. For these studies, cDCp were isolated from GFP trangenic mice and pulsed with dextran conjugated with Texas Red. DC division proceeded through several distinct stages (Fig. 5a and Movie S1).4 First, migration on the monolayer stopped. Second, the cells became spherical with loss of dendrites and veils; this shape change was the most reliable indicator for identifying DC that were about to divide. Third, the cells divided symmetrically, producing two daughters of equal size. Endosomes containing dextran were also distributed evenly to each daughter, consistent with our flow cytometry data in Fig. 4b. Fourth, the daughters remained in close contact for ~30–40 min. Finally, the daughters became increasingly motile, separated, and acquired characteristic DC morphology and migratory activity. DC replication resulted in the generation of network of interacting motile DC, each containing a similar amount of Ag (Fig. 5b and Movie S2).4


Figure 5
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  		FIGURE 5. cDCp create interacting DC networks. a, Live confocal microscopy of DC division. cDCp from GFP-transgenic mice were pulsed with dextran conjugated with Texas Red and cultured on S17. On the third day of culture, the cells were imaged by real-time confocal microscopy. Time course of DC division: upper panel, GFP; middle panel, dextran-Texas Red; lower panel, differential interference contrast. Original magnification, x1000. Also see Movie S1.4 b, DC colony that originated from a single dextran-pulsed cDCp. Left upper quandrant, merged; right upper quandrant, GFP; left lower quadrant, dextran-Texas Red; right lower quandrant, differential interference contrast. Original magnification, x600. Also see Movie S2.4

 
Spleen DC replication rate increases during adenovirus infection

It has been reported that DC turnover in lymphoid tissues increases during immune responses to antigenic and inflammatory stimuli (3, 29, 30). To determine whether the mechanism of Ag transfer by replicating DC is operational during an inflammatory process, mice were infected with Adv-βgal, which is highly immunogenic and rapidly induces an adaptive immune response (31). On day 0, 1, 3, or 6, these mice were injected with YG-latex beads and 8 h later were treated with BrdU for 12 h to identify dividing phagocytic DC (Fig. 6a). The rate of bead reuptake during the 12-h BrdU-labeling period was <2%, as determined by adoptively transferred DC. The number of spleen DC decreased by 50% on day 0, and was accompanied by a similar decrease in the proportion that incorporated BrdU (Fig. 6, b and c). Spleen DC numbers returned to baseline on day 1, and subsequently increased 3-fold on days 3 and 6 of infection. The rate of BrdU incorporation also rebounded, increasing 2-fold on day 1, 4-fold on day 3, and 5-fold on day 6, and was associated with an increased proportion of spleen DC in cell cycle (Fig. 6d). Bead phagocytosis and surface expression of MHC class II and CD86 by DC were enhanced on the first day of infection, and returned to control values by day 6 (Fig. 6, e–h). Remarkably, the proliferation rate of bead+ DC consistently paralleled that of bead– DC. β-galactosidase was detected in spleen cells at all time points (data not shown), which implied that viral-encoded Ags would be available to proliferating DC. We confirmed that an adaptive T cell immune response to adenovirus had developed by day 7 (data not shown). These results highlight that DC proliferation in spleen is dynamic, and that phagocytic DC continue to replicate and transmit internalized beads to progeny during an inflammatory process.


Figure 6
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  		FIGURE 6. DC replication increases during adenovirus infection. a, Experimental protocol. Mice were infected with 3 x 1010 virus particles of Adv-βgal. On the day indicated, mice were injected i.v. with YG-latex beads, and 8 h later were treated with BrdU for 12 h before cell recovery. b, Total number of conventional spleen DC on days 0, 1, 3, and 6 of infection. c, Percentage of bead+ and bead– spleen DC that incorporated BrdU over 12 h on days 0, 1, 3, and 6 after infection. d, Percentage of sorted spleen DC in the S/G2/M phase of the cell cycle. e, Frequency of DC that phagocytose beads on days 0, 3, and 6 of infection. f, Flow cytometric analysis of MHC class II and CD86 expression among gated CD11c+ lineage– DC on day 0. g, Fold increase of mean fluorescence intensity (MFI) of MHC class II expression by spleen DC as compared with control. h, Fold increase of MFI of CD86 expression by spleen DC as compared with control. Results are representative of three independent experiments. Error bars are mean ± SD.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Our study documents several previously unknown attributes of cDCp and DC. First, replication-competent cDCp can capture soluble Ag and particulates; second, cDCp generate proliferating DC that transmit internalized Ags to successive generations of their progeny during cell division; and third, Ag inherited by DC progeny is presented effectively to Ag-specific T cells (Fig. 7). Our novel findings have been corroborated by Liu et al. (32), who reported that MHC-peptide complexes can be detected on the surface of DC that have incorporated BrdU. Furthermore, we report for the first time that DC replication is augmented significantly during adenovirus infection, suggesting that this mechanism of Ag transfer may be a natural component of the response to infection and inflammation.


Figure 7
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  		FIGURE 7. Model of Ag transmission by replication-competent cDCp and DC.

Supplemental Movie Legends

Movie S1. DC Replication. Live imaging shows active cell division of a DC with transfer of intracellular dextran to progeny. Images were acquired every 10 s. Left upper quandrant (merged), right upper quandrant (GFP), left lower quandrant (dextran-Texas Red), right lower quandrant (DIC). Original magnification, x1200.

Movie S2. Replication-competent DC form network of motile Ag-bearing DC. cDCp from GFP-transgenic mice were pulsed with dextran conjugated with Texas Red and cultured on S17 for 3 days. Live imaging was performed on 3rd day of culture. Left upper quandrant (merged), right upper quandrant (GFP), left lower quadrant (dextran-Texas Red), right lower quandrant (DIC). Original magnification, x600.

 
cDCp are a distinct population of immediate DC precursors that exist in bone marrow and all lymphoid tissues (8). Evidence indicates that they are the chief source of conventional DC in spleen, at least during steady-state conditions (8, 9, 33). It has been suggested that cDCp might arise from a local progenitor population in spleen, but parabiotic studies by Liu et al. (32) indicate that they originate in blood, presumably migrating from bone marrow, where they outnumber mature DC by a 3:1 ratio (8). Adoptive transfer studies in our laboratory indicate that DC progeny of bone marrow and spleen cDCp are present for at least 7 days in spleen. Similarly, Naik et al. (9) found that the peak production of DC from adoptively transferred spleen cDCp was at 5 days, with some DC persisting for 8–10 days. These values are in agreement with those reported by Liu et al. (32) in parabionts. Thus, Ags internalized by cDCp have the potential to persist via their DC progeny for 10 days, which is significantly longer than the 2- to 3-day lifespan of DC predicted by previous BrdU incorporation studies.

The relative importance of the DC progeny of replicating Ag-bearing DC in activating T cells in vivo is unclear. The kinetics of T cell proliferation during immune responses would suggest that they have a minor role in initial T cell activation as compared with mature nondividing DC. Ag transmission by dividing DC, however, may enable Ag to persist in a population of DC despite rapid turnover of individual DC, and extend the duration of cognate interactions with T cells. It is interesting to note that DC expressing low density of MHC-peptide complexes on the surface seem to be important in the generation of memory T cell responses (34, 35).

We speculate that internalized Ags contained in endosomes and lysosomes are distributed randomly to each daughter during cell division, although the study by Liu et al. (32) suggests that MHC/peptide complexes located in the cell membrane might also be passed on to daughter DC. The inability of DC lysosomes to rapidly degrade internalized Ag most likely facilitates this process (11, 12). In the absence of a continuous supply of new Ag, two concurrent processes are envisaged to affect the amount of Ag in DC progeny: 1) the number of cell divisions (dilution); and 2) the rate of Ag degradation, which is influenced by lysosomal proteolytic activity and the susceptibility of the Ag to proteolysis (12). Previous studies of bone marrow-derived DC have shown that immature DC degrade internalized protein Ags slowly (36, 37). Because replicating DC are immature, based on their surface expression levels of MHC class II and costimulatory molecules and stimulatory capacity of allogeneic lymphocytes (16), the number of cell divisions may be the dominant factor that determines their Ag content.

Our finding that the in vitro and in vivo DC progeny of replicating OVA-pulsed cDCp could stimulate OT-1 T cells indicates that this mechanism of Ag transmission is functionally relevant. To ensure that there was no overlap between divided and undivided DC, we waited 3 days until several generations of divided DC were detectable, and then discarded the first generation of divided DC by FACS. Thus, the divided DC used in the T cell stimulation assays are comprised predominately of second and third generation progeny. The T cell proliferative response induced by equivalent numbers of divided DC was slightly lower than that by undivided DC. This difference might result from the lower levels of MHC class II expressed by divided DC as compared with undivided DC (data not shown). Alternatively, it might reflect differences in their Ag content.

cDCp cultured on a supportive stromal monolayer create networks of Ag-bearing DC with vigorous probing activity that resemble those visualized in lymph nodes by two-photon microscopy (38). It is hypothesized that such networks promote inter-DC Ag transfer and increase the available surface area of MHC-peptide complexes needed to stimulate T cells effectively. We found that Ag distribution to DC progeny was symmetrical, suggesting that each daughter of a divided DC should be equally effective in Ag presentation. Our coculture experiments with dextran- and OVA-pulsed cDCp also indicated that DC progeny acquired little or no Ag through inter-DC Ag transfer even though frequent interactions were clearly evident by live confocal microscopy (23, 24, 25). Whether the low level of DC apoptosis (data not shown) or the density of DC on the S17 monolayer accounts for the apparent absence of inter-DC Ag transfer is unclear.

Expansion of the number of DC in spleen has been observed in a variety of microbial infections, and has been attributed mostly to influx of new DC and increased DC survival (39, 40, 41, 42). Our findings with adenovirus infection indicate that increased DC proliferation is also an important factor. Increased replication of spleen DC could reflect influx of new replication-competent cDCp from the circulation, alteration of the cytokine milieu in spleen, or both. Whether the increased DC replication rate during infection involves those that have captured pathogen-derived Ags is unclear. In this regard, a previous report indicates that the nature of the phagocytosed cargo can influence DC function (43). It will also be important to investigate how cognate interactions with T cells influence DC proliferation. Finally, although we speculate that Ag transmitted through DC replication could enhance host defenses, it is noteworthy that this mechanism might also contribute to the replication and propagation of retroviruses in the immune system.


   Acknowledgments
 
We thank N. Iscove, M. Julius, R. Gorczynski, D. Grant, A. Kapus, and members of the Multiorgan Transplantation Program for helpful advice and critical reading of the manuscript; C. Cantin for FACS; and M. Shi for immunofluorescent microscopy.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
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 the Canadian Institutes for Health Research and Astellas (to M.S.C.). Presented in part at Immunology 2007, 94th Annual Meeting of the American Association of Immunologists, Miami Beach, FL, May 18, 2007. Back

2 Address correspondence and reprint requests to Dr. Mark S. Cattral, Toronto General Hospital, Robert McEwen Building, 11c-1247, 585 University Avenue, Toronto, Ontario, Canada M5G 2N2. E-mail address: mark.cattral{at}uhn.on.ca Back

3 Abbreviations used in this paper: DC, dendritic cell; Adv-βgal, adenovirus encoding bacterial β-galactosidase; cDCp, conventional DC precursors. Back

4 The online version of this article contains supplemental material. Back

Received for publication November 30, 2006. Accepted for publication June 12, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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