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In Vivo Analysis of Dendritic Cell Development and Homeostasis

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Science  17 Apr 2009:
Vol. 324, Issue 5925, pp. 392-397
DOI: 10.1126/science.1170540

Abstract

Dendritic cells (DCs) in lymphoid tissue arise from precursors that also produce monocytes and plasmacytoid DCs (pDCs). Where DC and monocyte lineage commitment occurs and the nature of the DC precursor that migrates from the bone marrow to peripheral lymphoid organs are unknown. We show that DC development progresses from the macrophage and DC precursor to common DC precursors that give rise to pDCs and classical spleen DCs (cDCs), but not monocytes, and finally to committed precursors of cDCs (pre-cDCs). Pre-cDCs enter lymph nodes through and migrate along high endothelial venules and later disperse and integrate into the DC network. Further cDC development involves cell division, which is controlled in part by regulatory T cells and fms-like tyrosine kinase receptor-3.

Dendritic cells (DCs) are immune cells that are specialized to capture, process, and present antigens to T lymphocytes in order to induce immunity or tolerance (1). Where commitment to DC development takes place, at what stage the monocyte lineage diverges from DCs, and the precise nature of the migrating DC precursor that moves from the bone marrow to the peripheral lymphoid organs are not known. These questions have been difficult to resolve in part because DC subsets are functionally and phenotypically diverse (2). For example, classical spleen DCs (cDCs) include two major functionally distinct subsets that are distinguished by the expression of a variety of C-type lectins and CD8 (24). Spleen and other tissues also contain plasmacytoid DCs (pDCs) that primarily initiate immune responses to nucleic acids (5, 6).

Lymphoid tissue cDCs, pDCs, and monocytes share a common progenitor called the macrophage and DC precursor (MDP) that is identified by its surface phenotype (Lin cKithi CD115+ CX3CR1+ Flt3+) (7, 8), whereas a distinct progenitor called the common DC precursor (CDP) (Lin cKitlo CD115+ Flt3+) is restricted to producing cDCs and pDCs (9, 10). Although monocytes can develop many of the phenotypic features of DCs under inflammatory conditions (1113), the cDC, pDC, and monocyte lineages separate by the time they reach tissues, and neither monocytes nor pDCs develop into cDCs under steady-state conditions (8, 14). Unlike monocytes and pDCs, cDCs in lymphoid tissues are thought to emerge from the bone marrow as immature cells that must further differentiate and divide in lymphoid organs (15, 16). Consistent with this idea, pre-cDCs that are restricted to the cDC lineage were isolated from the spleen and bone-marrow cultures containing fms-like tyrosine kinase receptor-3–ligand (Flt3-L) (17, 18). However, the relationship between MDP, CDP, and pre-cDC in vivo and the question of where the monocyte, pDC, and cDC lineages split have not been addressed (10, 14, 18).

We searched for MDPs and CDPs in the blood and spleen by means of flow cytometry but could only detect them in the bone marrow (Fig. 1A and fig. S1). Although pre-cDCs can be identified in the spleen by combining density centrifugation and flow cytometry (18), we speculated that these cells could be identified directly by expression of Flt3 and the chemokine receptor CX3CR1, which are expressed on other DC progenitors and also on mature cDCs (7, 10, 19). Indeed, we found a small but distinct population of lineage-negative CD11c+ major histocompatability complex (MHC) class II SIRP-αint Flt3+ cells (pre-cDCs) in the bone marrow (0.2%), blood (0.03%), spleen (0.05%), and lymph nodes (LNs) (0.03%) (Fig. 1B).

Fig. 1.

Isolation of pre-cDCs. (A) Presence of MDPs and CDPs in the bone marrow (BM), blood, or spleen (SP). NF, not found. (B) Identification of pre-cDCs in bone marrow, LNs, spleen, and blood. The bar graph shows the percentage of pre-cDC in each organ. 1 × 106 cells were acquired per sample. Lin-indicates cells that do not express CD3, CD19, Ter119, NK 1.1, or B220 antigens. (C) Donor-derived spleen cells (CD45.2+) were analyzed for cDC (CD11c+ MHC class II+), pDC (CD11cint B220+), and monocytes (CD11b+ CD11clo/–) 7 days after intravenous injection of 2 × 106 bone marrow cells or 1 × 105 pre-cDCs. (D) Analysis of donor-derived splenocytes after transfer with the indicated number of pre-cDCs from bone marrow, spleen, and blood. (E) Chimerism in parabiotic mice. In (B) and (C), the left side shows representative dot plots. The bar graphs summarize 2 to 4 independent experiments with 3 to 4 mice each. Error bars represent the mean ± SEM.

To determine whether pre-cDCs develop into cDCs in vivo, we compared them with unfractionated bone marrow cells in adoptive transfer experiments. After 7 days, unfractionated bone marrow gave rise to cDCs (10.2 ±1.2%), pDCs (1.9± 0.7%), and monocytes (13.5 ± 2.1%); in contrast, pre-cDCs from bone marrow, blood, or spleen yielded predominantly cDCs (88 to 95%) (Fig. 1, C and D, and fig. S2). These data indicate that pre-cDCs are indeed precursors to cDCs that arise in the bone marrow. They are restricted to the cDC lineage and have a reduced potential to produce monocytes or other lymphoid cells, including B, T, and natural killer (NK) cells (fig. S3). Pre-cDCs give rise to CD8 and CD8+ DCs but at slightly different ratios than are present in the unperturbed steady state (fig. S3).

Although the circulating pre-cDC had not been identified, parabiosis experiments suggested that they should be unequally exchanged between mice that share blood circulation because of their short half-life in the blood (15). To test this idea, we measured the exchange of cDCs and pre-cDCs in the bone marrow, blood, spleen, and LNs 50 weeks after parabiosis. We found incomplete (24%) chimerism of pre-cDCs in the blood and peripheral lymphoid organs even after prolonged parabiosis (Fig. 1E). We conclude that pre-cDCs arise from progenitors in the bone marrow and travel through the blood to peripheral lymphoid organs, where they further differentiate into cDCs.

To determine the relationship between pre-cDCs and other reported DC progenitors, we fractionated bone marrow cells from Cx3cr1gfp/+ reporter mice. In addition to MDPs (7) and CDPs (9, 10), we included an earlier developmental stage, myeloid progenitors (MPs) (Fig. 2A and fig. S2) (19). To determine the precursor-progeny relationship between MPs, MDPs, CDPs, and pre-cDCs, we performed adoptive transfer experiments in which purified progenitors were injected into the bone marrow of unirradiated 3- to 4-week-old recipient mice (8, 10). Two days after the transfer, MPs gave rise to MDPs, CDPs, and pre-cDCs, whereas MDPs only generated CDPs and pre-cDCs (Fig. 2B). Furthermore, at this early time point, CDPs produced only pre-cDCs (Fig. 2B). As previously reported, 7 days after transfer MDPs produced cDCs, pDCs, and monocytes, whereas CDPs gave rise to pDCs and cDCs but not monocytes (Fig. 2C and figs. S4 and S5) (810). Thus, MPs give rise to MDPs, which produce monocytes and CDPs. CDPs are further restricted in order to generate pre-cDCs and pDCs.

Fig. 2.

Progenitor-product relationship among MDP, CDP, and pre-cDC. (A) Phenotype of MP (Lin Sca-1 c-Kithigh CX3CR1), MDP (CX3CR1+ c-Kithi), and CDP (CD115+CX3CR1+c-Kitlo) in Cx3cr1gfp/+ mice. (B) Donor-derived bone marrow cells analyzed with flow cytometry for presence of MDPs, CDPs, and pre-cDCs 2 days after intra–bone marrow transfer of 5 × 104 MPs, MDPs, or CDPs. (C) Donor-derived splenocytes analyzed for the presence of cDCs, pDCs, and monocytes 7 days after transfer of MDPs or CDPs. (D) EGFP expression in indicated cell populations in LysM-CrexRosa26-StopfloxEGFP reporter mice (fig. S8). EGFP index is the percentage of EGFP+ cells divided by the percentage of EGFP+ Gr1high blood monocytes (20). In (B) and (C), the left side shows representative dot plots. The bar graphs summarize 2 to 4 independent experiments with 3 to 4 mice. Error bars represent the mean ± SEM.

Monocytes and pre-cDCs shared features with MDPs, but they diverged phenotypically from MDP in the bone marrow (figs. S6 and S7). To confirm the point of divergence between monocytes and cDCs, we made use of LysM-CrexRosa26-StopfloxEGFP mice (20). In these mice, monocytes and their progeny are irreversibly marked with enhanced green fluorescent protein (EGFP) expression because lysozyme promoter–driven Cre expression leads to deletion of the Stop sequence from Rosa26-StopfloxEGFP (Fig. 2D and fig. S8) (20). In contrast to bone marrow and blood monocytes and their progeny, MDP, CDP, pDCs, pre-cDCs, and cDCs did not express EGFP and therefore never went through a monocyte intermediate (Fig. 2D). We conclude that monocytes separate from the cDC lineage during the transition from MDP to CDP in the bone marrow, but our data do not rule out the possibility of an alternative monocyte-independent pathway of cDC development.

Similar to MDPs, monocytes and pre-cDCs in the bone marrow actively divided as measured by 4′,6′-diamidino-2-phenylindole staining and 5-bromo-2′-deoxyuridine incorporation; however, this was not observed in the spleen or in the blood (fig. S9). We conclude that pre-cDC expansion takes place primarily in the bone marrow and that upon differentiation into cDCs in lymphoid tissues, the latter undergo cell division (fig. S9) (17).

To examine the fate and distribution of pre-cDCs in lymphoid organs, we visualized them by means of two-photon laser scanning microscopy. Up to 5 hours after adoptive transfer, pre-cDCs were detected in close proximity to major blood vessels, primarily in the medullary side of the node and at the interface between the T cell zone and B cell follicles (Fig. 3A and movies S1 to S3; ratio of cortical to medullary pre-cDC = 0.41 at 2 to 5 hours after injection; n = 3 nodes). These vessels showed a cobblestone morphology and a distribution that is typical of high endothelial venules (HEVs) (movies S2 and S3). Consistent with the observation that pre-cDCs were frequently found near HEVs, pre-cDCs expressed CD62L, and antibody-mediated inhibition of CD62L prevented cDC accumulation in the LNs but not the spleen, in which pre-cDCs may enter the white pulp through the marginal sinuses (Fig. 3B and fig. S10) (21). Between 1 and 5 hours after injection, transferred pre-cDCs were sessile (average speed = 1.9 μm/min; SD = 1.0) and remained in contact with HEVs, exhibiting very little displacement (Fig. 3, C and D, and movies S2 and S3). After 16 to 18 hours, pre-cDCs and their progeny continued to localize to vessel-rich areas; however, they were no longer attached to blood vessels and displayed a more migratory behavior (average speed = 5.6 μm/min; SD = 3.0) (Fig. 3, C and D, and movies S4 and S5). In contrast, after 6 days the progeny of pre-cDCs were distributed throughout the LN paracortex and integrated into the endogenous DC network (Fig. 3A and movies S6 and S7; ratio of cortical to medullary pre-cDC = 1.95 at 6 days after injection; n = 3 nodes). These cells displayed dendritic morphology (Fig. 3A) and active probing behavior with very restricted displacement and reduced average speeds (3.7 μm/min; SD = 1.3) (Fig. 3, C and D, and movies S7 and S8) (22). We conclude that pre-cDCs enter the LNs through HEVs and then distribute themselves throughout the LN and assume typical DC behavior.

Fig. 3.

Multi-photon imaging of pre-cDCs. (A) Transferred cells in inguinal LNs 5 hours (top, green) and 6 days (bottom, blue) after pre-cDC transfer. Panels at right show morphology of transferred pre-cDCs (arrowheads). Scale bars, 1 mm (left) and 30 μm (right). (B) (Left) CD62L expression on pre-cDCs. (Right) Graph showing the number of LN cDCs or migratory DCs (mDCs) after treatment with antibody to CD62L. The graph represents the mean ± SD (n = 4 mice per group in 3 experiments). *P = 0.0286; **P < 0.0001; Student's t test. (C) Dynamic behavior, morphology, and position of pre-cDCs at indicated times. A cell track (yellow) is superimposed to visualize displacement. Scale bar, 50 μm. Far right shows superimposed two-dimensional (XY) tracks with starting coordinates set to the origin. The number of cells analyzed (n) is indicated. (D) Graphs show velocity and confinement of pre-cDCs at indicated times. Differences between columns are significant according to a Kruskal-Wallis test (a heterogeneity of speed, P = 0.0006; and meandering, P = 0.0055). *P < 0.05; **P < 0.01; ***P < 0.001; Dunn's multiple comparison test. All multi-photon data represent at least two independent experiments.

Depletion of CD4+Foxp3+ regulatory T cells (Tregs) leads to DC expansion, but how or where Tregs affect the DC developmental pathway has not been determined (23). To examine this, we investigated the effect of Treg depletion on DC development with the use of Foxp3DTR mice. These mice express the diphtheria toxin receptor (DTR) under the control of the Foxp3 promoter, and treatment of these mice with diphtheria toxin (DT) results in the specific deletion of Foxp3+ cells (23). We found no change in MDPs or bone-marrow pre-cDCs after Treg depletion. In contrast, spleen and LN pre-cDCs and cDCs increased by 2- and 12-fold, respectively (Fig. 4A) (23). In addition, the percentage of dividing cDCs in both spleen and LNs increased from 5 to 10%, which suggests that Treg depletion results in augmented pre-cDC and cDC division specifically in lymphoid organs (Fig. 4B).

Fig. 4.

Tregs control DC expansion in the peripheral lymphoid organs. (A) Numbers of MDP, pre-cDCs, and cDCs in the indicated organs after Treg depletion in Foxp3DTR mice. (B) Pre-cDC and cDC division in spleen and LNs at indicated time after Treg depletion. (C and D) Effect of the Flt3 inhibitor, Sutent, on Treg-depletion–induced cDC and B cell expansion. (E) Bar graph shows the relative percentage of Flt3–/– CD45.1 and Flt3+/+ CD45.2 DCs and B cells after a mixed bone-marrow transfer into FoxP3DTR CD45.1XCD45.2 mice after DT treatment. The panels represent two to four independent experiments. The bar graph represents the mean ± SEM (n = 3 to 4 mice).

Under steady-state conditions, the rate of cDC division in lymphoid organs is regulated in part by Flt3 receptor signaling (16). To determine whether the Flt3 pathway is required for Treg control of cDC expansion, we used Sutent (Pfizer, New York), a multi-targeted tyrosine kinase inhibitor with affinity for Flt3 (24). We found that Sutent treatment inhibited cDC expansion but not B cell expansion in Treg-depleted mice (Fig. 4, C and D). To further confirm the role of the Flt3 pathway in cDC expansion in response to Treg depletion, we adoptively transferred mixtures of bone marrow–derived DC progenitors from wild-type Flt3+/+ and Flt3–/– mice into Foxp3DTR recipients and measured their response to DT treatment. Whereas wild-type cDCs expanded, Flt3–/–- cDCs did not (Fig. 4E). We conclude that increased local production of Flt3-L by a yet-to-be-determined cell type is required for DC division in response to Treg depletion.

The precursor-progeny relationship of monocytes, cDCs, and pDCs has been debated since the discovery of DCs 35 years ago (1). Resolving this issue has been particularly difficult because monocytes can develop many of the phenotypic characteristics of DCs under conditions of inflammation in vivo or in the presence of certain cytokines in vitro (1113, 20). Furthermore, cDCs, pDCs, and monocytes share a common progenitor, the MDP (Fig. 2C). Several studies have shown that monocytes do not develop into cDCs, however, and that they make only a minor contribution to the lymphoid-organ DC network in the steady state (8, 25). We have defined the point of divergence between multipotential precursors and cDC restricted progenitors in the bone marrow and shown that the latter migrate through the blood to lymphoid tissues, where they and their progeny divide to fill the DC compartment. We further revealed that Tregs control DC development in the peripheral lymphoid organs in a Flt3-dependent manner (fig. S11). These newly recognized features of DC development in vivo open up the possibility of expanding or reducing DC numbers in vaccines and other clinical settings.

Supporting Online Material

www.sciencemag.org/cgi/content/full/1170540/DC1

Materials and Methods

Figs. S1 to S11

References

Movies S1 to S8

References and Notes

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