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*Stem Cells
The Journal of Immunology, 2006, 176: 7196-7206.
Copyright © 2006 by The American Association of Immunologists

In Situ Replication of Immediate Dendritic Cell (DC) Precursors Contributes to Conventional DC Homeostasis in Lymphoid Tissue1

Jun Diao*,¶, Erin Winter*,¶, Claude Cantin{dagger}, Wenhao Chen*,¶, Luoling Xu{ddagger}, David Kelvin{ddagger}, James Phillips§,|| and Mark S. Cattral2,*,¶

* Multiorgan Transplantation Program, Toronto General Research Institute, Toronto General Hospital, {dagger} Ontario Cancer Institute, {ddagger} Division of Experimental Therapeutics, § University Health Network and Hospital for Sick Children, Department of Surgery, and || Department of Pathology, University of Toronto, Toronto, Ontario, Canada


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The developmental biology of dendritic cells (DC) under physiological conditions remains unclear. In this study, we show that mouse CD11c+ MHC class II–lineage– cells are immediate precursors of conventional DC and are widely distributed in both bone marrow and lymphoid tissues. These precursors have a high clonal efficiency, and when cocultured on a supportive stromal monolayer or adoptively transferred in vivo, generate a population CD11c+MHC class II+ DC that retain limited proliferation capacity. During steady state conditions, a small proportion of immediate DC precursors (DCp) and DCs are dividing actively in bone marrow and spleen. Cytokines that initiate and support proliferation of immediate DCp were defined. Collectively, our findings provide evidence of a distinct development pathway for conventional DC in both bone marrow and lymphoid tissues and highlight the importance of in situ replication of immediate DCp and DC in maintaining conventional DC populations.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Dendritic cells (DC)3 in lymphoid tissues are in a dynamic balance. The estimated half-life of conventional mature DC in spleen during steady state conditions is only 2–3 days, and is reduced further by immune processes (1, 2). Although this rapid turnover of DC clearly mandates continuous replacement by a precursor population(s), the precise identity of immediate DC precursors (DCp) in lymphoid tissues remains uncertain (3). Circulating monocytes can differentiate into DC under defined cytokine conditions in vitro and in a variety of in vivo models (4, 5, 6, 7); however, evidence suggests their role in DC homeostasis in spleen is limited, at least during steady state conditions (8, 9). Reports of recent studies indicate that 4–8% of mature DC are actively replicating in the spleen during basal conditions, and this proportion increases during immune responses to antigenic stimuli (8, 10). In fact, mature CD11c+MHC class IIhigh DC were shown to expand 8- to 10-fold and generate CD11c+MHC class IIlow DC with regulatory properties. These observations have led to the novel concept that mature DC are not terminal cells, as previously thought, but are precursors for more DC (11). We identified a population of immediate DCp in bone marrow, which are CD11c+MHC class II– and do not stain for a variety of lineage markers including B220, CD3, CD19, DX5, GR-1, and F4/80 (12). A striking feature of this bone marrow population, which we refer to as B220–DCp, is its capacity to proliferate and generate a homogeneous population of conventional DC. Although a similar population of cells has been identified in the circulation (13, 14), its presence in secondary lymphoid tissues has not been established. Based on the known qualities of B220–DCp, we hypothesize that it may be an important precursor population for conventional DC in secondary lymphoid tissue.

To clarify the role of B220–DCp in DC development in lymphoid tissue, we examined its distribution in several secondary lymphoid organs. We also compared the phenotypic and proliferation properties of purified bone marrow and spleen B220–DCp as well as more mature DC populations. Our findings indicate that replication-competent B220–DCp are widely distributed in lymphoid tissue and contribute to DC homeostasis. Furthermore, we show that B220–DCp generates an intermediate population of CD11c+MHC class II+ DC, which retains a limited capacity to proliferate. Finally, we define cytokine conditions that promote and support proliferation of B220–DCp.


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

Male C57BL/6 and BALB/c mice were purchased from Charles River Laboratories. C57BL/6.SJL congenic and C57BL/6-Tg (ACTbEGFP)OPsb/J transgenic 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.

Abs and cytokines

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), CD16/32 (2.4G2) were purchased from BD Pharmingen. Anti-F4/80 (A3-1) was purchased from Serotec. These Abs were either unlabelled 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.

The following recombinant mouse cytokines were used: GM-CSF (BD Pharmingen); TNF-{alpha} (PeproTech); and IL-3, IL-11, stem cell factor (SCF), M-CSF, and Flt3 ligand (R&D Systems).

Cell isolation

Spleen, thymus, Peyer’s patches, and lymph nodes of 5–10 mice were minced, digested with collagenase and DNase for 0.5 h at 37°C, and incubated with EDTA. Mononuclear cells were isolated from these tissue preparations and bone marrow by Lympholyte-M (Cedarlane Laboratories) density gradient centrifugation (15), 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. For cells isolated from GFP transgenic mice, anti-I-Ab-biotin-allophycocyanin was used instead of anti-I-Ab-FITC. Lineage-negative CD11c+MHC II– cells (i.e., B220–DCp) 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.

Flow cytometry

Flow cytometry was performed on an Cytomics cytometer using Cytomics software (Beckman Coulter) as described previously (12). Briefly, cell suspensions were preincubated with anti-CD16/32 to block FcRs, then washed and incubated with the indicated mAb conjugates for 30 min at 4°C in a final volume of 100 µl of PBS containing 0.5% BSA and 2 mM EDTA. In all experiments, appropriate control isotype-matched mAbs were included to determine the level of background staining.

Cell culture

Sorted B220–DCp were cultured for 36 h in 96-well U-bottom culture plates at a cell density of 5 x 104/well in 200 µl of 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) in triplicate; in some wells LPS (1 µg/ml; Sigma-Aldrich) was added to the medium. In coculture experiments, sorted CD45.2+B220–DCp or CD45.2+CD11c+MHC class II+ DC (5 x 104) were incubated on a confluent monolayer of irradiated (25 Gy) stromal cells derived from S17 cells (a gift from K. Dorshkind, David Geffen School of Medicine at UCLA, Los Angeles, CA (16)). The cells were cultured in complete medium for 12–14 days with GM-CSF (1000 U/ml). Half the culture medium was replaced with fresh cytokine-containing medium every 3 days. Cultures were monitored by reverse lens microscopy (Nikon Eclipse TE 200) daily. TNF-{alpha} (100 U/ml) was added to some wells for the final 12–24 h of culture to induce DC maturation. At the end of the culture period, nonadherent cells were collected by gentle pipetting for further analyses. To quantify total cell expansion, cells adherent to the monolayer were also recovered after disrupting the monolayer with 0.25% trypsin and 1 mM EDTA. In other experiments, sorted cells were incubated in 96-well flat plates at a cell density of 1000 cells/well in complete medium containing single or various combinations of GM-CSF (1000 U/ml), M-CSF (10 ng/ml), IL-3 (10 ng/ml), IL-11 (10 ng/ml), SCF (10 ng/ml), and FLT3 ligand (100 ng/ml), at concentrations known to support DC development from early lymphoid and myeloid precursors (17). Every 3 days, half of the medium was replaced with fresh medium. After 7–8 days of culture, the cells were collected for further analysis.

CFSE labeling

Sorted cells were washed and resuspended in Hank’s solution at 105cells/ml, and incubated with 1 µM CFSE (Molecular Probes) for 15 min at 37°C. Cells were then washed first with RPMI 1640 containing 10% FBS then with RPMI 1640 without FBS twice, resuspended in complete medium, and cultured on S17 stromal monolayer for 3 days. To harvest cells, the monolayer was disrupted with 0.25% trypsin/1 mM EDTA and repeated pipetting. The recovered cells were washed, stained with anti-CD11c, anti-CD45.2, and anti-IAb mAbs, and analyzed by flow cytometry. Dead cells were excluded by propidium iodide (PI) staining. Analysis of cell division (CFSE fluorescence) was limited to CD45.2+ or CFSE+ cells. For in vivo studies, 106 cells/ml were incubated with 1 µM CFSE for 15 min at 37°C, washed once with RPMI 1640 containing 10% FBS then twice with HBSS; 5–10 x 105 CSFE-labeled cells were injected i.v. into congenic unirradiated or gamma-irradiated (6 Gy) recipients as described (18). The spleen was removed from recipients 3 or 7 days later; mononuclear cells were isolated by Lympholyte-M (Cedarlane Laboratories) density gradient centrifugation, stained with fluorochrome-conjugated Abs, and analyzed by flow cytometry.

Limiting dilution assay

Sorted GFP+B220–DCp were cultured by limiting dilution (an average 0.5, 1, 2, 4, 6, and 8 cells/well) on a S17 monolayer in 96-well plates with GM-CSF (1000 U/ml). Thirty wells containing an average of 1, 2, 4, 6, and 8 cells/well and 60 wells containing an average of 0.5 cells/well were established. Half the culture medium was replaced with fresh cytokine-containing medium every 3 days. Cell growth was monitored daily under an inverse fluorescence microscope. Wells containing colonies ≥6 cells on day 8 were counted as positive. Clonal efficiency was calculated by Poisson statistics.

Immunohistochemical staining

GFP+ B220–DCp cocultured on S17 cells for 8 days in 96-well plates were fixed with 4% paraformaldehyde for 10 min at room temperature. Cells were washed with TBS containing 10% FCS (washing buffer), and blocked with 10% mouse serum and 10 µg/ml anti-CD16/CD32 mAb at room temperature for 60 min. Cells were incubated with 100 µl of washing buffer containing 10 µg/ml biotin-conjugated anti-I-Ab or anti-CD11c Abs for 2 h at 4°C. Excess Ab was removed by washing before applying Texas Red streptavidin (Biomeda; 1 mg/ml diluted 1/250 with washing buffer) for 30 min at room temperature. Cells were visualized and photographed with an inverse florescence microscope (Nikon TE200) at a wavelength of 480 nm for GFP and 560 nm for Texas Red.

Electron microscopy

B220–DCp/S17 cell cocultures were mechanically removed from culture wells, fixed in 2.5% glutaraldehyde in PBS at 4°C for 2 h, washed in PBS, postfixed in 1% osmium tetroxide in PBS for 1 h, dehydrated in acetone and embedded in epon. Ultrathin sections were stained with lead citrate and uranyl acetate and analyzed with a Philips 400 electron microscope.

Determination of cell cycle status

FACS-purified bone marrow and spleen B220–DCp and CD11c+MHC class II+ DC pooled from three mice were fixed in 75% ethanol at 4°C for 16 h, and stained with PI (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).

BrdU staining

BrdU (1 mg; Sigma-Aldrich) was injected i.p. 3.5 h before recovery of bone marrow and spleen cells from groups of 5 mice. Cell sorting was performed to exclude Lin+ and autofluorescent cells. Purified CD11c+Lin– cells were stained with anti-I-Ab-PE, washed, resuspended in cold 0.15M NaCl, and fixed by dropwise addition of cold 95% ethanol. The samples were stored overnight at 4°C, stained with anti-BrdU Ab (BD Biosciences) as described by Kamath et al. (1), and analyzed by flow cytometry. Cells isolated from mice not injected with BrdU served as a control for background staining.

RT-PCR

Total RNA from 2.5x 105 sorted bone marrow B220–DCp and DC was extracted with TRIzol (Invitrogen Life Technologies) as per the manufacturer’s instructions. RNA was resuspended in RNase-free water and treated with DNase I (RNase-free; Invitrogen Life Technologies) to remove any contaminating genomic DNA. Before reverse transcription (RT), DNase I in samples was inactivated by addition of EDTA followed by incubation at 65°C for 10 min. Half of the treated RNA underwent first-strand cDNA synthesis using M-MuLV Reverse Transcriptase (MBI Fermentas), while the remaining RNA, to serve as a control, was subjected to identical conditions without RT. One microliter of the RT and control reaction products were used for the subsequent PCR. The sequences of the sense and antisense primers for mouse CCR1, CCR2, CCR5, CCR6, CCR7, CCR9, CXCR4, and beta-actin have been described previously (19). Samples were subjected to a total of 37 cycles using PCR conditions optimized for each primer set. The PCR products were subjected to electrophoresis on 1.5% agarose gel containing ethidium bromide and visualized by UV illumination. beta-actin was used as an internal control for RNA integrity. Nonreverse-transcribed RNA served as negative control.

Chemotaxis assays

Cell migration was determined in a microchemotaxis chamber (Neuroprobe) as described (20). B220–DCp were suspended at 1 x 106 cells/ml in migration medium (RPMI 1640 (without L-glutamine, 2-ME, or antibiotics) with 25 mM HEPES and 1% (w/v) BSA (Sigma-Aldrich)). CCL3 or CCL7 were dissolved in chemotaxis medium at graded concentrations and 25 µl was placed in the bottom chamber wells; 50 µl of cell suspension was placed in the upper compartment of the chamber. The chemotaxis chamber was assembled with a 0.5-µm pore size polycarbonate membrane and a silicon gasket separating the upper from the lower wells, and incubated in humidified air with 5% CO2 at 37°C for 1.5 h. At the end of the incubation period, membranes were removed, fixed, and stained with DiffQuick (Harlew). The number of migrated cells in three randomly selected high-power (x400) fields was counted by light microscopy. The results are expressed as mean (±SD) values of triplicate samples and are representative of at least three experiments performed.

Allogeneic MLR

Graded numbers of stimulator cells were seeded in triplicate in U-bottom 96-well culture plates (BD Biosciences). Responder spleen cells (1 x 105/well) from BALB/c mice were added to the wells in a total volume of 200 µl of RPMI 1640 complete medium, and cultured for 3 days in a humidified atmosphere of 5% CO2 in air at 37°C. The culture was pulsed with 1 µCi of [3H]thymidine (Amersham) 16 h before harvest, and collected onto glass fiber filters (Millipore); [3H]thymidine incorporation was quantified using a Beckman scintillation counter. Background controls with spleen cells or stimulator cells alone were included in all experiments and were always <500 cpm. Results are expressed as the mean cpm of triplicate cultures.

Statistics

Continuous variables are expressed as mean ± SD and were analyzed by the two-tail Student t test. A p value <0.05 is considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
B220–DCp are widely distributed in lymphoid tissue

CD11c+ mononuclear cells were enriched from bone marrow, spleen, thymus, Peyer’s patches, and lymph nodes with anti-CD11c magnetic beads, and analyzed by multicolor flow cytometry. Within the CD11c+ lineage (anti-CD3, CD19, B220, F4/80, and DX5) negative fraction, MHC class II expression ranged from undetectable to highly positive (Fig. 1a). A population of CD11c+MHC class II–lineage– cells, which we refer to as B220–DCp, comprise 62 ± 5% of total lineage–CD11c+ populations in bone marrow (n = 5 independent experiments), 19 ± 9% in spleen (n = 5), 30 ± 12% in thymus (n = 3), 17 ± 4% in Peyer’s patches (n = 3), and 9 ± 1% in lymph nodes (n = 3). B220–DCp from these different sites do not express the costimulatory molecules CD40, CD80, and CD86, and have similar expression levels of CD62L and CD44 (Fig. 1b). CD11b expression was slightly lower in bone marrow B220–DCp as compared with those in secondary lymphoid organs. From a single mouse, 5–10 x 104 B220–DCp were identified in bone marrow and spleen, and ≤1 x 103 cells from each of the other organs and tissues examined. It is possible that purified spleen B220–DCp may include some blood B220–DCp; however, when we isolated spleen B220–DCp after flushing the circulation with PBS, there was no difference in the number identified (data not shown).


Figure 1
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  		FIGURE 1. Distribution of B220–DCp. a, CD11c+ cells from the organs and tissues indicated were enriched by anti-CD11c magnetic beads and analyzed by multicolor flow cytometry. The gate used in the upper panel dot plots indicates cells that are CD11c+ and lineage (CD3, CD4, CD8{alpha}, CD19, B220, F4/80, DX5) negative. The numbers in lower panel dot plots indicate the percentage of CD11c+lineage– cells that are MHC class II– (i.e., B220–DCp). b, Expression of the indicated markers for bone marrow and spleen B220–DCp. Numbers in histograms indicate percentage of cells. Data are representative of three to five independent experiments. c, Surface expression of MHC class II and CD86 on sorted B220–DCp cultured in GM-CSF or GM-CSF and LPS at the indicated time points. d, Stimulation of allogeneic lymphocytes by freshly isolated spleen B220–DCp and spleen B220–DCp cultured in GM-CSF or GM-CSF and LPS for 36 h. Proliferation was assessed by [3H]thymidine incorporation; results are expressed as mean cpm x 103 ± SD. Data are representative of more than three independent experiments.

 
Further analysis of B220–DCp requires a stringent isolation protocol to minimize contamination by autofluorescent cells and other cell populations that express CD11c including NK cells and CD8+ T cells (21, 22). Because of the difficulty in isolating large numbers of purified B220–DCp from normal mice by FACS, in the subsequent studies we focused on bone marrow and splenic B220–DCp.

When cultured in medium containing GM-CSF, surface expression of MHC class II on sorted bone marrow and spleen B220–DCp was evident within 2 h, and expression of MHC class II and CD86 molecules progressively increased over 36 h (Fig. 1c). Most B220–DCp from both bone marrow and spleen expressed intracellular MHC class II, suggesting that the rapid appearance of MHC class II at the cell surface was recruited from pre-existing intracellular stores, as previously shown for immature DC (23, 24) (Fig. 1b). Exposure to LPS increased the level of expression of MHC class II and CD86, and increased their stimulatory capacity in allogeneic MLRs (Fig. 1d). These data indicate that B220–DCp in spleen and bone marrow have similar phenotypic and functional properties.

B220–DCp and DC are actively dividing in vivo

Recent studies indicate that mature DC are actively dividing in vivo (8, 10). To determine whether B220–DCp are dividing under physiologic conditions, we first determined their DNA content by PI labeling. We also examined conventional DC because previous studies of DC replication did not specifically exclude the B220+ plasmacytoid DC subset (10). To this end, we sorted all cells to high purity (Fig. 2a). Flow cytometric analysis of PI-labeled cells revealed that 9.4 ± 0.6% of bone marrow and 1.7 ± 1.1% of spleen B220–DCp were in the S/G2/M phases of the cell cycle, as compared with 3.6 ± 0.5% of bone marrow DC and 4.0 ± 1.4% of spleen DC (n = 6 experiments).


Figure 2
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  		FIGURE 2. B220–DCp and DC are actively replicating in vivo. a, Bone marrow and spleen B220–DCp and CD11c+MHC class II+ DC were sorted using the indicated gates and stained with PI; 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. The bar graph shows mean ± SD of six determinations with three mice per group. b, BrdU incorporation by bone marrow and spleen B220–DCp and DC. CD11c+lineage– cells were purified by FACS 3.5 h after BrdU injection, and analyzed by flow cytometry for BrdU and MHC class II staining; the dot plot quadrants were determined by isotype-matched controls (BrdU background staining in mice not treated with BrdU is <0.1%; data not shown) The bar graph shows mean ± SD of four determinations with three mice per group.

 
To further assess B220–DCp and DC proliferation, we measured BrdU incorporation 3.5 h after BrdU injection (Fig. 2b). CD11c+lineage– cells were purified by FACS and analyzed for BrdU and MHC class II expression by flow cytometry. BrdU was incorporated by 8 ± 2.1% of B220–DCp and 4.1 ± 0.2% of DC in bone marrow, and by 1.4 ± 0.4% of B220–DCp and 2 ± 0.3% of DC in spleen (n = 4 experiments).

Differential proliferation capacity of B220–DCp and DC

Although these data indicate that both B220–DCp and DC are actively dividing in vivo, the proliferation capacity and relationship between these populations was unclear. We reported previously that stromal monolayers of S17 cells can support proliferation of bone marrow B220–DCp (12). Similarly, Zhang et al. (10) has shown that stromal cells derived from neonatal spleen can support proliferation of mature CD11c+MHC class IIhigh DC. Therefore, we compared the proliferation capacity of sorted B220–DCp and CD11c+MHC class II+ DC subpopulations from bone marrow and spleen in our coculture system.

B220–DCp from both bone marrow and spleen produced multicellular clusters on the monolayer that increased in number and size over 10 days; by 12 days, large numbers of single cells were released spontaneously into the culture medium. As compared with the input number of cells, the total number of cells recovered per culture of bone marrow and spleen B220–DCp increased 70 ± 22- and 19 ± 5-fold, respectively. In contrast, bone marrow and spleen CD11c+MHC class II+ DC did not expand (Fig. 3a). We wondered whether the poor proliferation of mature DC in this coculture system was specific to the use of S17 cells. However, when the same cells were cultured on monolayers of other stromal cell lines (OP9, NIH 3T3) or on primary stromal cells derived from neonatal skin fibroblasts and spleen, similar results were obtained (data not shown).


Figure 3
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  		FIGURE 3. Differential proliferation capacity of B220–DCp and DC. Bone marrow and spleen B220–DCp and CD11c+MHC class II+ DC were cocultured on S17 monolayers in the presence of GM-CSF (1000 U/ml) for 12 days. TNF-{alpha} (100 U/ml) was added to some wells for the final 24 h of culture. The progeny were recovered at 12 days for further analysis. a, Mean ± SD fold increase in cell number as compared with the initial number of cells placed in culture. b, Cell surface expression of the indicated markers by B220–DCp progeny. c, Stimulation capacity of allogeneic lymphocytes. A total of 105 BALB/c splenic lymphocytes was stimulated by graded numbers of B220–DCp progeny. Results are expressed as mean cpm x 103 ± SD. d, Morphology of Giemsa-stained progeny of spleen B220–DCp before and after stimulation with TNF-{alpha} (original magnification, x40). e, Cell surface expression of the indicated markers by spleen B220–DCp progeny before (thin line) and after TNF-{alpha} stimulation (dark line). Isotype control is indicated by dotted line. Data are representative of more than three independent experiments.

 
There was no discernable difference in the progeny of bone marrow and splenic B220–DCp. All were homogeneous morphologically, displayed immunophenotypic and functional characteristics of immature conventional DC (i.e., high expression levels of CD11c and MHC class II, low to moderate levels of CD40, CD80, and CD86, and potent stimulation of allogeneic lymphocytes), and could be matured further by exposure to TNF-{alpha} or LPS (Fig. 3, b–e).

To examine the possibility that mature DC were proliferating at an earlier time point, we examined CFSE-labeled populations 2 and 3 days after culture. However, so few cells were recovered after culture of bone marrow and spleen CD11c+MHC class II+ DC, they could not be analyzed further. By contrast, analysis of B220–DCp from bone marrow and spleen revealed that by the third day of culture 80–90% of the recovered cells represented the progeny of the starting population, with most completing ≥2 division cycles (Fig. 4a). MHC class II expression was up-regulated on the proliferating progeny of B220–DCp.


Figure 4
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  		FIGURE 4. B220–DCp generate proliferating CD11c+MHC class II+ DC. a, B220–DCp from BM and spleen were labeled with CFSE and cocultured on S17 stromal cells in the presence of exogenous GM-CSF, and recovered 3 days later for analysis of CD11c, MHC class II, and CFSE expression. Cell proliferation is measured by loss of 50% fluorescence intensity with each division, as indicated. b, Adoptive transfer of bone marrow and spleen B220–DCp. A total of 5 x 105 B220–DCp from CD45.2 mice was injected i.v. into unirradiated CD45.1 congenic mice. Mice transplanted with CD45.1 bone marrow only were used as a negative control (data not shown). At day 3, splenocytes were isolated and expression of CD45.2 or CFSE was used to identify donor-type cells. Cell proliferation is measured by loss of 50% fluorescence intensity with each division, as indicated. Data is representative of three independent experiments. c, A total of 5 x 105 bone marrow and spleen CD11c+ B220–DCp from CD45.2 mice was transplanted into irradiated (6 Gy) CD45.1 congenic mice, along with 5 x 104 CD45.1 bone marrow cells. At day 7, splenocytes were isolated and expression of CD11c and CFSE on CD45.1+ cells was determined by flow cytometry. Data are representative of three independent experiments.

 
To clarify whether B220–DCp and DC could expand in vivo, sorted bone marrow and spleen B220–DCp and DC from CD45.2 mice were labeled with CFSE and injected i.v. into nonirradiated CD45.1 mice. At 3 days, the mean percentage of donor-derived cells recovered in the spleen per 5 x 105 cells injected was 10.5 ± 4.9% (n = 6) for bone marrow B220–DCp and 3.7 ± 2.4% (n = 3) for spleen B220–DCp; over 50% of these cells had divided at least once, as determined by CFSE intensity. Most of the divided and undivided cells expressed MHC class II (Fig. 4b). By contrast, <0.1% of injected bone marrow or spleen CD11c+MHC class II+ DC could be recovered in the spleen (data not shown). Bone marrow and spleen B220–DCp and/or their progeny, but not mature DC, were also detected in low numbers (200–300) in lung, lymph nodes, Peyer’s patches, and bone marrow 3 days after injection. The poor recovery of mature DC is consistent with our in vitro findings and adoptive transfer experiments reported by others (25, 26, 27), and indicates that mature DC have limited potential to expand as compared with B220–DCp.

Flow cytometric analysis of B220–DCp division by CFSE staining beyond 3 days was difficult in normal mice because of the low ratio of donor-to-recipient-derived cells. Therefore, we injected bone marrow and spleen B220–DCp i.v. into sublethally (6 Gy) irradiated CD45.1+ congenic recipients and recovered their spleen 7 days later (12) (Fig. 4c). We found that most bone marrow and spleen B220–DCp had divided by this time point, and many of their progeny had completed multiple division cycles.

Clonal efficiency of B220–DCp

It was not clear why splenic B220–DCp had less capacity to expand than bone marrow B220–DCp. The CFSE data indicated that the rate of proliferation of dividing bone marrow and spleen B220–DCp during the first 3 days was similar. Therefore, we speculated that there might be a difference in the proportion of cells capable of dividing. To address this possibility, we cultured the cells by limiting dilution to determine their clonal efficiency. Purified bone marrow and spleen B220–DCp from GFP transgenic mice were used for these studies, which enabled us to visualize clone development from single cells on the stromal monolayer. Clones of GFP+ cells emerged from single cells within 3–5 days and many continued to enlarge with some containing up to 80 cells at 8 days (Fig. 5a). Most cells in the clusters had dendritic processes and were MHC class II+ (Fig. 5b) Clonal expansion decreased at 8–10 days and stopped by 12 days (data not shown), suggesting that the progeny have a finite proliferative capacity. Clonal efficiency for bone marrow and splenic B220–DCp was 52.6 ± 21.3 and 9.8 ± 4.2% (n = 4), respectively (Fig. 5c). The lower clonal efficiency of spleen B220–DCp would account, at least partly, for the reduced number of progeny recovered after in vitro coculture or adoptive transfer as compared with bone marrow B220–DCp. These data, nonetheless, establish that replication-competent B220–DCp are present in both bone marrow and spleen.


Figure 5
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  		FIGURE 5. Clonal efficiency of B220–DCp. a, Fluorescence microscopy shows clone development from single B220–GFP+ DCp sequentially photographed over 8 days. Original magnification, x20. b, Detection of MHC class II by immunofluorescence on day 8. Original magnification, x40. c, Limiting dilution analysis of bone marrow and spleen B220–DCp. Purified GFP+B220–DCp were deposited by limiting dilution (average of 0.5, 1, 2, 4, 6, and 8 cells/well) onto the S17 monolayer in 96-well plates with GM-CSF (1000 U/ml) and cultured for 8 days. Wells with clones containing more than six cells were deemed positive. Each data point is the mean ± SD of four independent experiments.

 
Chemokine receptor expression of B220–DCp

To gain insight into the mechanisms that govern migration of B220–DCp, we examined the expression of mRNA encoding for chemokine receptors in bone marrow B220–DCp by RT-PCR. Bone marrow B220–DCp express CCR1, CCR2, CCR5, CCR9, and CXCR4 (Fig. 6a), and in contrast with mature bone marrow DC do not express CCR7. In chemotaxis assays, we found that CCL3, which binds to CCR1 and CCR5, induces B220–DCp migration in a dose-dependent manner, whereas CCL7, which binds to CCR1 and CCR2, had no effect on migration (Fig. 6b).


Figure 6
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  		FIGURE 6. B220–DCp migrate in response to CCL3. a, RT-PCR analysis of mRNA for chemokine receptors on sorted bone marrow B220–DCp (DCp) and DC (DCm). b, Chemotactic response of sorted bone marrow B220–DCp to graded concentrations of CCL3 and CCL7. Control is medium alone. Results are mean ± SD (*, p < 0.01 compared with control). Data are representative of three independent experiments.

 
Critical signals that support B220–DCp proliferation

The signals that promote and support replication of B220–DCp are unknown. Because S17 cells could stimulate B220–DCp proliferation, we initiated investigations to identify growth factors that might be involved in this process. In Transwell experiments, we established that S17 supernatant could stimulate bone marrow B220–DCp proliferation (Fig. 7a). GM-CSF alone did not stimulate B220–DCp proliferation, but was strikingly synergistic with S17 supernatant, increasing B220–DCp expansion from 6.1- to 18.7-fold after 8 days of culture. Because the range of growth factors produced by S17 cells is large, we decided to limit our investigation to six cytokines (GM-CSF, Flt3 ligand, IL-3, M-CSF, IL-11, and SCF) based on their high mRNA expression in S17, as determined by RT-PCR (data not shown), and previous evidence implicating their involvement in DC ontogeny. In isolation, none of these cytokines supported proliferation of B220–DCp, although all slightly improved survival (data not shown). When combined as a mixture, however, 6.8-fold expansion occurred by 8 days (Fig. 7b). Withdrawal of each cytokine individually from the mixture revealed that GM-CSF, Flt3 ligand, and M-CSF were the most important. We next examined these three cytokines individually and in various combinations to determine their relative importance (Fig. 7c). No cell expansion occurred unless both GM-CSF and Flt3 ligand were present; addition of M-CSF to this combination increased cell proliferation slightly (3.4- vs 2.5-fold, respectively). The progeny displayed morphological and immunophenotypic characteristics of DC, regardless of the cytokine conditions (Fig. 7d).


Figure 7
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  		FIGURE 7. GM-CSF and Flt3 ligand in combination support B220–DCp proliferation. a, Bone marrow B220–DCp were deposited in the upper chamber of the Transwell with or without a S17 monolayer in the lower chamber or placed directly on the monolayer, and cultured for 8 days. GM-CSF was added to some wells. b, Bone marrow B220–DCp were cultured for 8 days in medium containing a mixture of GM-CSF, Flt3 ligand, M-CSF, IL-3, SCF, and IL-11. Single cytokines were withdrawn from the mixture to determine the relative importance of each cytokine. c, Bone marrow B220–DCp were cultured for 8 days with GM-CSF, Flt3 ligand, and M-CSF individually and in various combinations. The mean ± SD fold increase in cell number as compared with the initial number of cells placed in culture is shown in the bar graphs. d, Flow cytometric analysis shows expression of indicated markers of recovered cells that had been cultured in GM-CSF and Flt3 ligand or GM-CSF, Flt3 ligand, and M-CSF for 8 days. e, Transmission electron micrographs of cultured B220–DCp on stromal monolayer (original magnification, x6000 on left, and x8000 on right). A direct contact point between a B220–DCp and stromal cell is indicated by the arrow. Data are representative of three independent experiments.

 
Direct contact with S17 monolayers is more effective than S17 supernatant in expanding B220–DCp (Fig. 7a). Fibronectin, a matrix protein produced by stromal cells, has been reported to promote proliferation of mature DC (10). When B220–DCp were cultured on fibronectin-coated or -uncoated plates in the presence of S17 supernatant and GM-CSF, there was no difference in the number of cells recovered (data not shown). Using transmission electron microscopy, we detected distinct contact points between B220–DCp and stromal cells (Fig. 7e), suggesting that intercellular signaling may be important for survival and proliferation of B220–DCp and their progeny.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Our characterization of B220–DCp strongly supports their involvement in DC homeostasis in bone marrow and lymphoid tissue. As compared with mature DC, B220–DCp have a significantly higher capacity to replicate. We also found that B220–DCp generate an intermediate population of CD11c+MHC class II+ cells in vitro and after adoptive transfer, which supports the notion that actively dividing DC in vivo arise from B220–DCp rather than mature DC (Fig. 8). Mouse spleen contains three populations of conventional DC that are distinguishable by cell surface expression of CD4 and CD8{alpha} (28). Although adoptively transferred bone marrow B220–DCp generate both CD8{alpha}+ and CD8{alpha}– DC subsets in spleen (12), we cannot rule out the existence of other distinct precursors for each DC subset as suggested by previous kinetic studies (25).


Figure 8
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  		FIGURE 8. Model of conventional DC development.

 
The traditional view of DC ontogeny holds that DC populations in lymphoid tissue and peripheral organs come from migrating precursors that subsequently differentiate into mature DC (29). Our finding of an extensive distribution of B220–DCp with a substantial proportion of B220–DCp and DC actively dividing supports an alternative model in which DC arise from proliferating resident immediate precursors in situ. Replication-competent B220–DCp and DC have also been detected in inflamed tissues and organs suggesting that in situ DC proliferation may be more pervasive than expected (30, 31). The ability to augment DC expansion locally may be an important mechanism for rapidly amplifying the number of DC available for Ag processing and presentation.

The composite characteristics of DC-restricted differentiation and the high capacity to replicate distinguish B220–DCp from other cell populations known to generate DC including pluripotent stem cells, monocytes, and B220+ plasmacytoid-DCp (6, 7, 32). Based on Poisson statistics, we show that bone marrow B220–DCp can proliferate with essentially absolute efficiency. The lower clonal efficiency of spleen B220–DCp as compared with bone marrow B220–DCp may reflect an intrinsic difference between these purified populations. It is also conceivable that splenic B220–DCp are comprised of more than one cell population with variable capacities to proliferate even though we have been unable to detect any consistent phenotypic or functional differences in the two populations so far. Alternatively, the splenic microenvironment may alter the clonal capacity of B220–DCp. Current studies are aimed at addressing which of these explanations is the most likely.

MHC class II expression is up-regulated on the proliferating progeny of cultured and transplanted bone marrow and spleen B220–DCp. By contrast, we found that B220–DCp predominate over mature DC in bone marrow despite having a 2- to 3-fold higher proliferation rate than DC in vivo. We hypothesize that the bone marrow microenvironment favors B220–DCp proliferation without differentiation, and thereby permits maintenance of a reservoir of replication-competent precursors ready to be released into the circulation in response to stress and inflammatory stimuli. Indeed, studies have shown that proinflammatory mediators including TNF-{alpha} and MIP-1{alpha} can rapidly mobilize B220–DCp into the circulation (30, 33). Our finding that bone marrow B220–DCp migrate in response to CCL3 (MIP-1{alpha}) supports the hypothesis that bone marrow is a major source of these circulating precursors. Studies in parabiotic mice and other evidence, however, suggest that some proliferating spleen DC could also arise from resident progenitors in spleen (8, 34).

Although our data indicate that DC are actively proliferating in vivo, it not possible to determine based on BrdU or PI labeling whether all DC are proliferating or are capable of proliferating. In fact, quite to the contrary, freshly isolated CD11c+MHC class II+ DC expand poorly in vitro and after adoptive transfer as compared with the MHC class II+ progeny of B220–DCp. Several explanations for this apparent paradox can be envisioned. First, CD11c+MHC II+ DC in vivo are heterogeneous with respect to maturation level and proliferation potential; even actively dividing cells have completed a variable number of division cycles since the immediate precursor stage. In this regard, our data indicates that the proliferation capacity of the progeny of B220–DCp wanes with successive divisions and is ultimately lost. We believe the categorical terminology used to describe DC (e.g., mature, immature) belies the continuous evolution of various biologic attributes, including proliferation capacity, that occurs as precursors progress toward terminally differentiated mature DC (Fig. 8). Second, some mature CD11c+MHC class II+ cells in vivo may arise from precursors with limited proliferation capacity such as monocytes. Finally, it is possible that mature DC are more sensitive to the isolation technique than B220–DCp and undergo terminal maturation and apoptosis shortly after culture. This explanation seems unlikely, however, because CD11c+MHC class II+ DC purified from collagenase digests of B220–DCp/stromal cocultures can still proliferate when replated on new stromal monolayers (data not shown).

We found that proliferating CD11c+MHC class II+ cells arising from B220–DCp in bone marrow and spleen produce conventional DC with potent immunostimulatory capacity. By contrast, Zhang et al. (10) reported that mature CD11c+MHC class IIhigh DC expand and generate regulatory DC when cocultured on spleen stromal monolayers. Whether differences in the experimental conditions used in our and their study accounts for the variation in DC generated merits further investigation.

Our analysis of the conditions that support proliferation of B220–DCp demonstrates the importance of both the stromal microenvironment and cytokine milieu in this process. The beneficial effect of the stromal monolayer on B220–DCp proliferation can be replaced partly by a mixture of soluble cytokines, and of those examined, the combination of Flt3 ligand and GM-CSF was critical. The importance of GM-CSF is not surprising as its role in various aspects of DC development is well documented. The function of Flt3 ligand in DC ontogeny beyond primitive hemopoietic progenitors (e.g., common myeloid and lymphoid precursors) has been controversial, however, because Flt3 receptor expression is low or undetectable on circulating immediate DCp and mature DC (12, 35, 36). Interestingly, the addition of M-CSF to the culture medium slightly increased B220–DCp proliferation and did not promote macrophage differentiation, as has been shown for monocytes (37, 38). The ability to expand immediate DCp in liquid medium will facilitate future studies. Direct contact with S17 cells was more efficacious than S17 supernatant alone in promoting B220–DCp expansion. Preliminary studies suggest that direct contact with the monolayer improves survival of B220–DCp immediately after culture, which increases the number of cells available to proliferate (data not shown). Our electron microscopy studies show intimate contacts form between B220–DCp and stromal cells; how this augments cell expansion or survival remains unclear, however. Fibronectin has been reported to induce proliferation of mature DC (10). Although fibronectin had no effect in our system, it is possible that other matrix proteins such as heparin sulfate proteoglycans, which are produced by S17 cells and can modulate GM-CSF activity, are involved (39, 40).

Collectively, the results presented here support a role for B220–DCp in the development of conventional DC in bone marrow and spleen. Their presence in other secondary lymphoid organs suggests they are also involved in maintaining DC in those organs as well. Further studies are required to determine whether B220–DCp are absolutely required or whether they represent one of several pathways for conventional DC development. Active proliferation of immediate DCp and DC in secondary lymphoid organs raises several intriguing questions. Does DC proliferation simply serve to maintain DC populations that are rapidly turning over, or is it involved somehow in Ag processing and dissemination? What initiates, regulates, and stops DCp proliferation? Answers to these questions should provide important insights into DC biology.


   Acknowledgments
 
We thank N. Iscove for advice and R. Gorczynski for critical reading of the manuscript.


   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 (to M.S.C.), National Institutes of Health (A155024-01; to M.S.C.), and Physicians Services Foundation (to M.S.C.). 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; DCp, DC precursor; SCF, stem cell factor; PI, propidium iodide; RT, reverse transcription. Back

Received for publication September 30, 2005. Accepted for publication March 24, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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