CCR2⁺CD103⁻ intestinal dendritic cells develop from DC committed precursors and induce interleukin 17 production by T cells

Abstract

The identification of intestinal macrophages (mϕs) and dendritic cells (DCs) is a matter of intense debate. While CD103⁺ mononuclear phagocytes (MPs) appear to be genuine DCs, the nature and origins of CD103⁻ MPs remain controversial. We show here that intestinal CD103⁻CD11b⁺ MPs can be separated clearly into DCs and mϕs based on phenotype, gene profile and kinetics. CD64-CD103⁻CD11b⁺ MPs are classical DCs, being derived from Flt3L-dependent, DC-committed precursors, not Ly6C^hi monocytes. Surprisingly, a significant proportion of these CD103⁻CD11b⁺ DCs expresses CCR2 and there is a selective decrease in CD103⁻CD11b⁺ DCs in mice lacking this chemokine receptor. CCR2⁺CD103⁻ DCs are present in both the murine and human intestine, drive IL-17a production by T cells in vitro, and show constitutive expression of IL-12/IL-23p40. These data highlight the heterogeneity of intestinal DCs and reveal a bona fide population of CCR2⁺ DC, which is involved in priming mucosal Th17 responses.

Introduction

The intestine is exposed constantly to many antigens and innate immune stimuli, including dietary constituents, commensal bacteria and pathogens ¹. The intestinal immune system must discriminate between these different agents, mounting protective immunity against pathogens, but developing active tolerance to harmless materials. If this process fails, inappropriate responses can lead to inflammatory bowel diseases and celiac disease. Mononuclear phagocytes (MPs) in the lamina propria (LP) such as dendritic cells (DCs) and macrophages (mϕs) are central to these events, serving distinct, yet complementary functions. Whereas DCs migrate to draining lymph nodes and prime naïve T cells ^2-5, mϕs are sessile phagocytes that scavenge bacteria and damaged cells. They also maintain the expansion of antigen-specific regulatory T cells through the production of IL10 ^6,7 and promote epithelial barrier integrity ⁸.

Because of their distinct functions, it is likely that mucosal DCs and mϕs play different roles in disease, meaning they need to be characterised as precisely as possible. However this has been the source of much confusion and controversy ^4,9-14, largely because many of the phenotypic markers used to discriminate DCs and mϕs are insufficiently specific. Initial studies defined intestinal DCs simply on the basis of CD11c and MHCII co-expression, but most if not all mϕ in the mucosa also express these markers ^{3,10,13,15,16}. More recent work has used CD103, CD11b and CX3CR1 expression to identify three major populations of CD11c⁺MHCII⁺ MPs. Two of these express CD103, lack CX3CR1 and are either CD11b⁺ or CD11b^{− 3,5,17,18}. Based on their derivation from DC-committed precursors (pre-DCs) and genetic profiles, it is generally agreed that these CD103⁺ MPs are bona fide DCs ^11,19,20. The third population of CD103⁻CD11b⁺ MPs is less well understood. Although they express CX3CR1 ^5,16,21 and were originally considered to be monocyte-derived DCs (mo-DCs) ^19,20, recent transcriptional analyses suggest they are more similar to mϕ than DCs ^9,11. However we recently identified CD103⁻CD11b⁺CX3CR1^int cells migrating in pseudo-afferent lymph that are bona fide CD103⁻ DCs, based on their responsiveness to Flt3L and lack of mϕ markers such as F4/80 or CD64 ⁵ and our unpublished observations). Although analogous cells have been described in steady state LP ^5,16,22,23, confusion remains over the relative contribution of DCs and mϕs to the CD103⁻CD11b⁺ LP population and their developmental origin remains contentious. In previous work, we found that Ly6C^hi monocytes could not generate CD103⁺ or CD103⁻ subsets of bona fide DCs in steady state colon ^16,24. However a more recent study suggested that CD103⁻ DCs in the mucosa are indeed monocyte-derived, on the basis that their accumulation and/or development involves a CCR2 dependent precursor ²³.

Here we identify a population of genuine CD103⁻ DCs in the LP that are phenotypically, genetically and kinetically distinct from mϕs. Like their CD103⁺ counterparts, these CD103⁻ DCs arise from Flt3L dependent, DC-committed precursors and not from Ly6c^hi monocytes. Significantly we also demonstrate the presence of CCR2-expressing CD103⁻CD11b⁺ DCs in the murine and human LP, which have a selective ability to prime IL-17a-producing CD4⁺ T cells in vitro. The existence of this novel subset of bona fide DCs may help explain previous conflicting results and provides insights into functional compartmentalisation among mucosal DC populations.

Results

Expression of F4/80 and CD64 defines two distinct CD103⁻CD11b⁺ mononuclear phagocyte populations in intestinal lamina propria

To begin to determine the origins of CD103⁻ MPs in the small intestinal LP, we first set out to establish a gating strategy that would allow accurate discrimination between CD103⁻ DCs and mϕs. After first identifying intestinal MPs as CD11c⁺MHCII⁺ cells amongst live leukocytes, we found 3 discrete populations based on CD103 and CD11b expression; a majority population of CD103⁻CD11b⁺ cells, together with smaller numbers of CD103⁺CD11b⁺ and CD103⁺CD11b⁻ cells (Figure 1a). A small population of CD103⁻CD11b⁻ cells was also identified, but as these may derive from isolated lymphoid follicles, rather than the LP itself ⁵, we did not examine them further. To distinguish between DCs and mϕs, MPs were next examined for expression of the pan-mϕ marker F4/80, together with CD64, which has recently been described to be specific for mϕs in the intestine ^16,24. Consistent with the general consensus that CD103⁺ MPs are classical DCs ^3,4,18-20,25, none of the CD103⁺ MPs expressed either F4/80 or CD64 (Figure 1a). In contrast, the CD11c⁺MHCII⁺CD103⁻CD11b⁺ cells were heterogeneous. While ~85% of this population were F4/80⁺CD64⁺ mϕs, the remainder were entirely negative for both CD64 and F4/80, suggesting they may be DCs (Figure 1a,b). Identical subsets of CD64⁺ and CD64⁻ MPs were present in steady state colonic LP, albeit in different proportions, with the CD103⁺CD11b⁻ and CD103⁻CD11b⁺ subsets both outnumbering the CD103⁺CD11b⁺ DCs in this tissue (Supplementary Figure 1a-c). Interestingly, there was an inverse correlation between the proportions of CD103⁺CD11b⁺ DCs and CD64⁺ mϕs throughout the length of the small and large intestine, whereas the proportions of the other DC subsets were not significantly different between the various segments of the small intestine, before increasing in the colon (Supplementary Figure 1b, c). All CD64⁻ subsets could also be identified amongst migratory CD11c⁺MHCII^hi MPs in the steady state MLN, whereas CD64⁺ MPs were virtually absent (Supplementary Figure 1d,e).

Figure 1. CD64⁻CD103⁻CD11b⁺ intestinal MPs are bona fide DCs.

(a) Intestinal MPs were defined as CD11c⁺MHCII⁺ cells amongst live CD45⁺ lamina propria cells, analysed for CD103 and CD11b expression and the resulting populations of MP were examined for CD64 and F4/80 expression. (b) Proportions of MP populations amongst total CD11c⁺MHCII⁺ cells. Data are representative of at least 15 independent experiments with n=3/4 per experiment. (c) SI LP MP populations were FACS-purified and Zbtb46 expression assessed by real-time PCR. The results shown are expressed relative to HPRT using the 2^−ΔΔct method with mϕs set to 1 and are pooled from 2 independent experiments each with cells pooled from 9-12 mice on each occasion. (d) Total SI LP digests were incubated at 37°C or 4°C with pHrodo E. coli bioparticles for 15 minutes and assessed for phagocytic ability. Bar chart shows mean ΔMFI (MFI 37°C-MFI 4°C) + 1SD, n=4. ***p<0.001 vs mϕs, one way Anova with Bonferroni post test. (e) Expression of CD272, CD26, MerTK, CD14, and CX3CR1 on SI LP MP populations (coloured) compared with appropriate background controls (shaded grey). (f) Populations of SI LP MP as a proportion of total live leukocytes in WT and Flt3L^−/− mice. Data are pooled from 2 independent experiments ***p<0.005, Student’s t test. (g,h) Mice received one injection of 1mg/ml BrdU i.p., followed by 0.8mg/ml BrdU in the drinking water for 3 days, before being returned to normal drinking water. Mice were culled immediately following cessation of BrdU feeding, or 3 or 9 days later and MP subsets identified in SI LP (g) and amongst migratory DC in the MLN (h). Data are the proportions of BrdU⁺ cells amongst each subset and are representative of 2 independent experiments with n=3/4 per time point. Black asterisks represent significant differences between D3 and D3+9 for all DC populations, while coloured asterisks represent significant differences between D3 and D3+3 in MLN. ***p<0.001, **P<0.01, *p<0.05 Student’s t test.

CD64⁻CD103⁻CD11b⁺ MPs are bona fide DCs

To clarify further the nature of the CD103⁻CD11b⁺ cells, we examined other features that define DCs and mϕs. Unlike the CD64⁺ mϕs, all subsets of CD64⁻ MPs, including the CD103⁻CD11b⁺ cells, expressed mRNA for the DC-specific transcription factor Zbtb46 ^26,27 (Figure 1c) and were not phagocytic as assessed by the ability to take up bacterial particles into acidified vesicles (Figure 1d). Whereas all the CD64⁺F4/80⁺CD103⁻CD11b⁺ mϕσ expressed the mϕ-specific markers MerTK and CD14 ^9,11, the CD64⁻F4/80⁻CD103⁻CD11b⁺ MPs lacked expression of these markers, as did the prototypic CD103⁺ DCs. Conversely, CD64⁻F4/80⁻CD103⁻CD11b⁺ MPs expressed the recently defined DC-specific markers CD26 and CD272, while CD64⁺F4/80⁺CD103⁻CD11b⁺ MPs did not (Figure 1e). Consistent with previous studies ^{3,16,18,21,24,28}, CX3CR1 was expressed at high or intermediate levels on CD64⁺F4/80⁺CD103⁻CD11b⁺ mϕs, but was absent from CD103⁺ DCs (Figure 1e). However, the CD64⁻CD103⁻CD11b⁺ MPs within the mucosa all expressed intermediate levels of CX3CR1 (Figure 1e), consistent with the CD103⁻CD11b⁺ DCs we identified recently in pseudo-afferent intestinal lymph ⁵.

Finally we examined the dependence of the different MP subsets on the DC-specific growth factor Flt3L ^29,30. As expected, normal numbers of CD11c⁺CD64⁺ mϕs were present in the intestine of Flt3L^−/− mice (Figure 1f). In contrast, all CD11c⁺CD64⁻ subsets including the CD64⁻CD103⁻CD11b⁺ MPs were dramatically reduced in the LP of Flt3L^−/− mice compared with WT controls (Figure 1f). Taken together, these data confirm the presence of bona fide CD103⁻CD11b⁺ DCs in the steady state LP.

Intestinal DCs divide in situ and turn over rapidly in the steady state

Previous work has shown that intestinal mϕs and CD103⁺ DC have distinct population dynamics in vivo ^3,18. However, these studies considered all CD103⁻CD11b⁺ MPs to be mϕs. Having now identified bona fide CD103⁻CD11b⁺ DCs in LP, we used BrdU pulsing to determine how their population dynamics compared with mϕs and CD103⁺ DCs in vivo.

To examine this, BrdU was administered continuously for 3 days and its incorporation was assessed immediately after its withdrawal and again 9 days thereafter. After 3 days of BrdU feeding, a substantial majority (70-85%) of both subsets of CD103⁺ DCs was BrdU⁺, whereas only 15-25% of CD64⁺ mϕs had been labelled at this time (Figure 1g). Importantly CD103⁻CD11b⁺ DCs displayed BrdU-labelling identical to the CD103⁺ DC subsets and very distinct from CD64⁺ mϕs (Figure 1g). 9 days after BrdU withdrawal, the proportion of BrdU⁺ cells amongst all of the CD64⁻ DC subsets had decreased to less than 10%, while the proportion of BrdU⁺ CD64⁺ mϕs remained steady, indicating relative longevity compared with DCs (Figure 1g). In contrast to LP, where maximal BrdU labeling was achieved upon BrdU withdrawal at day 3, only 30-50% of the CD64⁻CD103⁺ and CD103⁻ populations amongst the CD11c⁺MHCII^hi migratory cells in MLN were BrdU⁺ at this time (Figure 1h). The frequency of BrdU⁺ cells amongst migratory DCs in the MLN increased further 72hr after BrdU withdrawal, consistent with migration of BrdU⁺ cells from the LP. Consistent with the short lifespan of DCs, only 15-20% of MLN DCs remained BrdU⁺ 9 days after BrdU⁺ withdrawal. Resident CD11c⁺MHCII^int DCs in MLN had a distinct BrdU profile, achieving maximal labeling before withdrawal of BrdU (data not shown).

CD103⁻CD11b⁺ intestinal DCs arise from committed DC precursors

Previous studies that did not take the heterogeneity of CD103⁻CD11b⁺ MPs into account suggested that they were derived exclusively from Ly6C^hi monocytes ^19,20,31. Thus we next investigated the origin of the bona fide CD103⁻CD11b⁺ DCs, first by adoptive transfer of Ly6C^hi monocytes from CX3CR1^+/gfp mice into monocytopenic CCR2^−/− mice (Supplementary Figure 2a,b). 5 days after transfer, donor Ly6C^hi monocytes and their progeny could be identified in the SI LP by their expression of CD45.1 and CX3CR1-GFP. At this time all donor cells were CD11c⁺MHCII⁺F4/80⁺, with none being found within the CD11c⁺F4/80⁻ DC gate (Figure 2a). Furthermore no progeny of transferred Ly6C^hi monocytes could be found in the MLN (data not shown). Thus in our hands, Ly6C^hi monocytes cannot act as progenitors for rigorously defined CD103⁻ DCs in the steady state intestine and nor do monocyte-derived MP migrate to the MLN from the LP under these conditions.

Figure 2. CD64⁻CD103⁻CD11b⁺ DCs arise from DC committed precursors.

(a) 1×10⁶ Ly6C^hi monocytes sorted from the BM of CX3CR1^+/GFP CD45.1⁺× CD45.2⁺ mice were transferred into resting CCR2^−/− CD45.2⁺ recipients. 5 days later, donor-derived cells were identified amongst live SI LP cells as CD45.1⁺CX3CR1-GFP⁺ and examined for expression of CD11c, MHCII and F4/80. Data are representative of two recipient mice from a single experiment. (b) CD45.2⁺ WT mice were injected with 2×10⁶ Flt3L secreting B16 tumour cells. 10-14 days later pre-DC were FACS-sorted, labeled with Cell Trace Violet proliferation dye and transferred i.v. into CD45.1⁺ congenic mice. 5 days later, donor-derived cells were identified amongst live SI LP cells as CD45.2⁺CD45.1⁻ Cell Trace Violet⁺ and examined for expression of CD11c, MHCII and F4/80. (c) Representative CD103 and CD11b staining by CD11c⁺MHCII⁺ donor cells in the SI LP compared with endogenous CD11c^hiMHCII⁺ cells. (d) Scatter plot shows proportion of CD11c⁺MHCII⁺ donor cells in each population defined by CD103 and CD11b expression as gated in C. Data are pooled from 3 independent experiments with n=10.

Given their independence from monocytes, we next tested whether CD103⁻CD11b⁺ DCs were derived from committed DC precursors (pre-DCs) ^32-34, as has been shown for mucosal CD103⁺ DCs ^19,20. Pre-DCs from BM (Lin⁻CD11c^intCCR9⁻B220⁻CD135⁺SIRPα^intCD117⁻; Supplementary Figure 2b,c) were FACS-purified from CD45.2⁺ mice which had received B16 tumours secreting Flt3L 10 days earlier, labelled with CellTrace Violet dye and transferred into unmanipulated, steady state, congenic CD45.1⁺ recipients. Donor derived cells could already be found in the LP 24 hours after transfer, at which time most had begun to upregulate both MHCII and CD11c (Supplementary Figure 2d,e). The expression of both these markers then increased progressively until their levels became equivalent to those on endogenous DCs. By 5 days after transfer, most donor cells had acquired MHCII expression and upregulated CD11c, but remained F4/80⁻, consistent with them being DCs (Figure 2b and Supplementary Figure 2d,e). The transferred pre-DCs not only gave rise to both CD103⁺CD11b⁻ and CD103⁺CD11b⁺ DC populations in LP, but there was also a clear population of CD103⁻CD11b⁺ donor-derived cells that lacked F4/80 expression, demonstrating directly that pre-DC can give rise to CD103⁻CD11b⁺ DCs (Figure 2c,d). Donor pre-DCs also gave rise to all three DC populations amongst CD11c⁺MHCII^hi migratory DCs in the MLN, as well as in the colonic LP (data not shown).

CD103⁻CD11b⁺ DCs are partially dependent on CCR2

Our results indicate that CD64⁻CD103⁻CD11b⁺ intestinal MPs are DCs and are distinct from monocyte derived mϕs. However a recent study has identified a similar population of CX3CR1^intCD11c⁺MHCII⁺ MPs in inflamed mucosa which were characterised as monocyte-derived DCs on the basis of their expression of Zbtb46 and CCR2 dependence ²³. On this basis it was concluded that some CD103⁻ DCs differentiate from monocytes, which require CCR2 for their egress from the BM ³⁵. Therefore, we examined whether deletion of CCR2 affected the abundance of our DC subsets in the steady state mucosa. Compared with WT controls, CCR2^−/− mice had normal proportions and absolute numbers of CD103⁺ DCs. However they had a significant reduction in CD64⁻CD103⁻CD11b⁺ DCs in the SI LP, and this was mirrored by a reduction in the CD103⁻ subset of CD11c⁺MHCII^hi migratory DCs in the MLN (Figure 3a,b). A small reduction in the number of CD103⁺CD11b⁻ DCs was also observed amongst migratory DCs in the MLN (Figure 3b). The selective CCR2 dependence of CD103⁻CD11b⁺ DCs and CD64⁺ mϕs was cell intrinsic, as it was replicated when these populations were derived from CCR2^−/− precursors in CD45.1 WT:CD45.2 CCR2^−/− mixed bone marrow chimeras (Figure 3c). Whereas WT and CCR2^−/− BM had a roughly equivalent ability to populate the CD103⁺ subsets of DCs, comparable with CCR2-independent eosinophils, the repopulation of CD103⁻CD11b⁺ DCs was biased towards WT BM, with a WT:CCR2^−/− ratio of ~3 (Figure 3c,d). Similar results were observed for CD103⁻CD11b⁺ DCs amongst migratory DCs in the MLN of chimeric mice (Figure 3d). Consistent with their dependence on replenishment by CCR2-dependent Ly6C^hi monocytes in the steady state ^16,24, there was a substantial reduction in the numbers of CD64⁺ mϕs in CCR2^−/− LP and virtually all mϕs in the LP of the mixed chimeras were of WT origin, similar to the pattern seen with blood Ly6C^hi monocytes (Figures 3a,d).

Figure 3. Partial dependence of CD103⁻CD11b⁺ LP DCs on CCR2.

(a) SI LP DCs from CCR2^−/− and WT mice were examined for expression of CD103 and CD11b, with the numbers representing the frequency of each population as a percentage of total CD11c⁺MHCII⁺CD64⁻ DCs. Scatter plot shows absolute numbers of each MP population in SI LP of CCR2^−/− and WT mice. (b) Migratory MLN DCs from CCR2^−/− and WT mice were examined for expression of CD103 and CD11b, with the numbers representing the frequency of each population as a percentage of total mig. DCs. Scatter plot shows absolute numbers of each mig. DC population in the MLNs of CCR2^−/− and WT mice. Data are representative of two independent experiments with n=3/4 per experiment *p<0.05, **p<0.005, Student’s t test. (c) CD45.1⁺/CD45.2⁺ WT mice were lethally irradiated and reconstituted with a 50:50 mix of BM from CD45.1⁺ WT and CD45.2⁺ CCR2^−/− mice. Chimerism amongst the populations of CD11c⁺MHCII⁺ MPs from SI LP was assessed 8 weeks later. (d) Ratio of CD45.1 WT:CD45.2 CCR2^−/− derived cells amongst CD11c⁺MHCII⁺ MPs and eosinophils from SI LP, blood Ly6C^hi monocytes (mo), and migratory DCs in the MLN of chimeric mice. ***p<0.001 vs mϕs, one way ANOVA with Bonferroni post-test. Data are representative of 2 independent experiments with n=3 per experiment.

Two populations of CD103⁻CD11b⁺ intestinal DCs can be defined by CCR2 expression

We reasoned that this partial dependence on CCR2 could reflect heterogeneity amongst the CD103⁻CD11b⁺ subset of DCs and therefore analysed CCR2 expression by the DC subsets in SI LP. Whereas CD103⁺ DCs were mainly CCR2⁻, there were two clear populations of CD103⁻CD11b⁺ DCs, the majority (~75-85%) being CCR2⁻ and a smaller, but distinct fraction being CCR2⁺ (Figure 4a). Because of this heterogeneity, we thought it important to examine if pre-DCs could give rise to both CCR2⁻ and CCR2⁺ CD103⁻CD11b⁺ DCs, which had been investigated as a single population in our earlier analyses. As with the endogenous population, CD103⁻CD11b⁺ DCs derived from transferred pre-DCs were heterogeneous for CCR2 expression 5 days after transfer, confirming that the CCR2⁺ DCs were derived from pre-DCs (Figure 4b). FACS-purified CCR2⁺CD103⁻CD11b⁺ DCs also expressed mRNA for Zbtb46 and Flt3 at levels equivalent to CD103⁺ DCs and their CCR2⁻CD103⁻ DC counterparts, confirming their DC lineage (Figure 4c). Furthermore, both the CCR2⁻ and CCR2⁺ subsets of CD103⁻CD11b⁺ DCs in the LP expressed mRNA for CCR7 at similar levels to CD103⁺ DCs, showing they have the potential to migrate out of the mucosa to the MLN (Figure 4c). Consistent with this, CCR2⁺CD103⁻CD11b⁺ DCs could be identified amongst migratory DCs in the steady state MLN (Figure 4d).

Figure 4. CCR2 expression defines two populations of intestinal CD103⁻CD11b⁺ DCs.

(a) Representative CCR2 staining on CD11c⁺MHCII⁺CD64⁻ DC populations from SI LP of WT mice, compared with total DCs from CCR2^−/− mice as a control. The scatter plot shows the proportion of CCR2⁺ cells in each SI LP DC population as a percentage of total CD11c⁺MHCII⁺CD64⁻ cells, with each point representing a mouse and the horizontal line representing the mean. Data are representative of 3 independent experiments with n=4 per experiment. (b) Representative CCR2 staining on CD103⁻CD11b⁺ DCs amongst endogenous (CD11c⁺MHCII⁺CD64⁻) and donor-derived (Violet⁺, CD45.2⁺, CD11c⁺, MHCII⁺) cells in the SI LP 5 days after transfer of CD45.2⁺ pre-DCs into unmanipulated CD45.1⁺ WT recipients. (c) Q-PCR analysis of mRNA for Zbtb46, Flt3 and Ccr7 by CCR2⁺ and CCR2⁻ subsets of CD103⁻CD11b⁺ DCs FACS-purified from the SI LP of WT mice, compared with CD103⁺ DC subsets and CD64⁺ mϕs. Data are from a single experiment with 10 mice pooled and are expressed relative to HPRT using the 2^−ΔΔct method with mϕs set to 1. (d) Representative CCR2 staining on subsets of DCs amongst CD11c⁺MHCII^hi migratory DCs in wild type MLN, and in total migratory DCs from CCR2^−/− mice used as a control. Scatter plot shows the proportion of CCR2⁺ cells in each SI LP DC population amongst total CD11c⁺MHCII⁺CD64⁻ cells, with each point representing a mouse and the horizontal line representing the mean. Data are representative of 2 independent experiments with n=4 per experiment.

CCR2⁺ and CCR2⁻ CD103⁻CD11b⁺ intestinal DCs are functionally distinct

Having identified two distinct populations of CD103⁻CD11b⁺ DCs based on their expression of CCR2, we next sought to determine the functional consequences of this heterogeneity. Thus SI LP DC subsets were FACS-purified, pulsed with OVA protein and co-cultured with CFSE-labelled, OVA-specific TcR transgenic naïve CD4⁺ T cells. All DC subsets, including the CCR2⁺ and CCR2⁻ populations of CD103⁻CD11b⁺ DCs induced similar levels of OTII T cell proliferation, as assessed by CFSE dilution (Figure 5a-c). The two subsets of CD103⁺ DCs generated T_Reg in vitro as determined by intracellular expression of FoxP3 and this appeared to be somewhat more efficient than either of the CCR2⁺ and CCR2⁻ subsets of CD103⁻CD11b⁺ DCs that had similar activity (Figure 5a). CD103⁺CD11b⁻ DCs induced the differentiation of considerably more IFNγ producing CD4⁺ T cells than all other DC subsets; there were no differences between the CD103⁺CD11b⁺ DCs and the CCR2⁺ or CCR2⁻ subsets of CD103⁻CD11b⁺ DCs in this respect (Figure 5b). However the CCR2⁺CD103⁻CD11b⁺ DCs were more effective than the other subsets at driving the differentiation of T_h17 cells in vitro (Figure 5c). Notably, this property within the CD103⁻CD11b⁺ DCs was masked when they were compared with other subsets as a single population (data not shown).

Figure 5. CCR2-expressing CD103⁻CD11b⁺ intestinal DCs drive T_h17 responses.

3×10⁴ FACS-purified DC subsets from SI LP were pulsed with 2mg/ml OVA protein and co-cultured for 4 days with 1×10⁵ FACS-purified naïve CFSE labeled OTII T cells. FoxP3 (a) IFNγ (b) and IL-17a (c) expression by live T cells was assessed by intracellular staining. Scatter plots show fold change in FoxP3 (a) IFNγ (b) and IL-17a (c) production induced by each DC population compared with that induced by CCR2⁻CD103⁻CD11b⁺ DCs. Mean values (+1SD) for the CCR2⁻ DCs were 0.341% (0.181), 1.051% (0.798) and 0.291% (0.09) amongst total CD4⁺ T cells for FoxP3, IFNγ and IL17a expression respectively. Data are shown as mean ±1SD pooled from three independent experiments. *p<0.05, **p<0.01, ***p<0.001, one way ANOVA with Bonferroni post-test. (d,e) Whole SI LP digests were incubated for 4.5 hours in the presence (+LPS) or absence (unstimulated) of 100ng/ml LPS with brefeldin A and monensin. Scatter plot shows percentage of IL-6⁺ (d) or IL-12/IL-23p40⁺ cells (e) amongst CCR2⁻ or CCR2⁺CD103⁻CD11b⁺ DCs assessed by intracellular cytokine staining. *p<0.05, **p<0.01, ***p<0.001 vs CCR2⁺CD103⁻CD11b⁺ DCs. Data are representative of 1 or 2 independent experiments with n=3/4 per experiment.

Efficient generation of T_h17 cells by the CCR2⁺ subset of CD103⁻CD11b⁺ DCs is associated with higher frequency of IL-12/IL23p40 production

In an attempt to understand why CCR2⁺CD103⁻CD11b⁺ DCs were effective in priming T_h17 cells, we examined their production of T_h17 polarising cytokines. Intracellular cytokine staining showed that there were ample proportions of IL-6 producing cells amongst all DC subsets under steady state conditions and if anything, thise were slightly lower for the CCR2⁺CD103⁻CD11b⁺ DCs than the CCR2⁻CD103⁻CD11b⁺ or CD103⁺CD11b⁺ subsets (Figure 5d and data not shown). However the CCR2⁺CD103⁻CD11b⁺ population was the only one in SI LP to contain IL-12/IL-23p40 producing DCs under steady state conditions, as well as showing the most robust increase in the frequency of IL-12/IL-23p40 producing cells in response to TLR4 stimulation. Under these conditions, ~30% IL-12/IL-23p40⁺ DCs were found within this subset, compared with ~10% in other subsets (Figure 5e). TLR4 stimulation had no effect on the already high frequencies of IL-6 producing cells amongst any subset (Figure 5d). Given that the CCR2-expressing CD103⁻CD11b⁺ DCs did not show any difference in their ability to prime T_h1 responses, these findings indicate that their preferential effect on T_h17 cells may be explained most readily by an enhanced ability to produce IL-23.

CCR2⁺CD103⁻CD11b⁺ LP DCs both express and are dependent on the transcription factor IRF4

A defining feature of the CD11b⁺ lineage of conventional DCs in mice is dependence on the IRF4 transcription factor for their development ^22,25,36-39, and we and others have shown recently that lack of IRF4 expression in CD11c⁺ cells leads to a selective defect in CD103⁺CD11b⁺ intestinal DCs that correlates with impaired T_h17 responses in vivo ^22,36. As these results appear to contrast with our discovery of a CCR2⁺ subset of CD103⁻CD11b⁺ DCs that is more efficient in inducing T_h17 cell differentiation than the other subsets, we thought it important to assess how IRF4 might control their development. Like CD103⁺CD11b⁺ DCs in LP, both the CCR2⁺ and CCR2⁻ subsets of CD103⁻CD11b⁺ DCs expressed IRF4, but not IRF8 (Figure 6a). Conversely and as expected, the CD103⁺CD11b⁻ DCs showed the opposite pattern, expressing IRF8, but not IRF4 (Figure 6a). As we found previously, CD11c-cre × IRF4^fl/+ mice had a substantial decrease in the numbers and proportions of CD103⁺CD11b⁺ DCs in SI LP and interestingly, these animals also showed a significant defect in the CCR2⁺ subset of CD103⁻CD11b⁺ DC (Figure 6b). In contrast, CD103⁺CD11b⁻ and CCR2⁻CD103⁻CD11b⁺ DCs were unaffected in CD11c-cre × IRF4^fl/+ LP. Thus the loss of CCR2⁺CD103⁻CD11b⁺ DCs may contribute to the defective T_h17 cell generation in these mice.

Figure 6. CCR2⁺CD103⁻CD11b⁺ DCs express IRF4 and require it for their existence in the LP.

(a) DCs in the SI LP were examined for their intracellular expression of IRF4 and IRF8 and compared with isotype controls. (b) Representative FACS plots and mean proportions and numbers of DC subsets in the SI LP of CD11c-cre⁻ × IRF4^Fl/+ (CRE⁻) or CD11c-cre⁺ × IRF4^Fl/+ (CRE⁺) mice. *p<0.05, **p<0.01, ***p<0.001 Student’s t test. Data are representative of 2-3 independent experiments with n=3-6 per experiment.

Putative CCR2-expressing CD103⁻ DC equivalents in the human intestine

Having identified CCR2⁺CD103⁻CD11b⁺ DCs in the murine intestine, we next sought to determine if equivalent cells were present in the human intestinal LP. By excluding CD14⁺CD64⁺ cells which represent mϕs ²⁴, DCs were readily identifiable in human colonic LP amongst live CD45⁺CD11c⁺HLA-DR⁺ cells (Figure 7). As in mouse LP, examination of CD103 and SIRPα expression identified 3 distinct populations of human DC populations and the CD103⁻SIRPα⁺ DCs were heterogeneous for CCR2 expression (Figure 7). Thus the CCR2 based heterogeneity of CD103⁻ mucosal DCs is maintained across species.

Figure 7. Putative CCR2⁺CD103⁻CD11b⁺ DC equivalents exist in healthy human colonic lamina propria.

DCs were identified in healthy human colon as live, CD45⁺, CD14⁻, CD64⁻, CD11c^+, HLA-DR⁺ and expression of CD103 and SIRPα was assessed. CD103⁻SIRPα⁺ DCs were then examined for CCR2 expression compared with CCR2 isotype control. Data are representative of 2 independent experiments.

Discussion

The nature and origins of intestinal DCs have been contentious for several years. Although the presence of two CD103⁺ DC subsets based on CD11b expression has been recognised for some time and they are generally accepted to be conventional DCs ^{3,17,19,20,40-42}, the existence of genuine CD103⁻ DCs amongst CD11c⁺MHCII⁺ MPs has been less certain. Initially considered to be pro-inflammatory “DCs” ^19,20,43, later studies defined CD103⁻CD11b⁺ MPs as mϕs, on the assumption that CD103 and CX3CR1 were mutually exclusive markers of DCs and mϕs respectively ³. Here we demonstrate that more rigorous strategies reveal heterogeneity within this population in the steady state intestine. In addition to confirming recent findings that the majority of CD103⁻CD11b⁺ express the mϕ markers CD64 and F4/80 ^16,22,24,44, we show definitively that a population of CD64⁻CD103⁻CD11b⁺ DCs is present in steady state LP. A similar subset can be found in pseudo-afferent intestinal lymph ⁴⁴. Like their CD103⁺ counterparts, the CD103⁻CD11b⁺ DCs in LP expressed Zbtb46 ^26,27, as well as CD26 and CD272, markers identified by the Immunological Genome Consortium as being specific to DCs ¹¹. Conversely they lacked the mϕ-restricted markers CD14 and MerTK ⁹, and showed poor phagocytic activity compared with CD64⁺ mϕs. Importantly, the CD64⁻CD103⁻ DCs expressed Flt3 and were virtually absent in the LP of Flt3L^−/− mice. The CD103⁻CD11b⁺ DCs also had a short half-life in vivo and showed evidence of proliferation in situ, properties identical to DCs in other peripheral tissues ²⁵ and to their CD103⁺ counterparts in the SI LP. In stark contrast, CD64⁺CD103⁻CD11b⁺ mϕs were long-lived and non-cycling.

Our studies using adoptively transferred precursors underlined the heterogeneity of intestinal CD103⁻ MPs, by demonstrating that CD103⁻CD11b⁺ intestinal DCs derived from DC-committed precursors. This contrasts with previous reports which concluded that CD103⁻ “DCs” were derived from Ly6C^hi monocytes. However these studies used less precise phenotyping strategies and using intense depletion of endogenous myeloid cells, which may alter the fate of donor cells and their progeny ^19-21,23. In our hands, Ly6C^hi monocytes were unable to generate CD103⁻CD11b⁺ DCs and gave rise only to F4/80⁺ mϕs in healthy mucosa. Because we were limited by the numbers of monocytes available for transfer, we cannot exclude the possibility that some genuine DCs might arise if there is expanded entry of monocytes into the mucosa. However our studies suggest that this is likely to be a rare event under steady state conditions. In contrast pre-DCs could clearly give rise to all populations of DCs, including the novel CCR2⁺CD103⁻CD11b⁺ subset we identified. Whether these distinct subsets arise from the same or multiple subsets of these progenitors remains unclear. A recent study has suggested that β7 integrin-expressing DC precursors had a preferential ability to repopulate mucosal CD103⁺ DCs ⁴⁵ but, it is currently unclear how these “pre-mucosal” DCs relate to the pre-DCs used here and in other studies ^34,45. Although our transferred pre-DCs lacked the B220 expression reported on the “pre-mucosal” DCs, a proportion were β7⁺ (our unpublished observations) and therefore some progenitors with selective gut-homing properties may be present in our populations. How these β7⁺ precursors specifically contribute to intestinal DC populations remains to be investigated. The idea that there might be independent precursors of each DC subset could explain why the proportions of donor-derived DCs in the intestine did not always mimic those found amongst host DCs at different times after transfer, as each lineage may develop at different rates. Alternatively, these time dependent differences might reflect developmental relationships within the same lineage and these issues will be the subject of future investigations.

At first sight, our conclusions that all mucosal DCs derive from pre-DCs appear to contradict a recent study by Zigmond et al, in which a population of mucosal Ly6C^loCD11b⁺Zbtb46⁺ cells, likely to comprise our CD103⁻ DCs, was depleted by in vivo administration of anti-CCR2 antibody ²³. As a result, it was concluded that these DCs were derived from CCR2-dependent Ly6C^hi monocytes. An explanation for this apparent discrepancy comes from our discovery of a population of pre-DC derived, CCR2-expressing CD103⁻CD11b⁺ DCs that have all the properties of the other bona fide DC in LP and are found amongst migratory DC populations in the MLN. Importantly, CCR2-deficient mice displayed a selective reduction in the numbers of CD103⁻CD11b⁺ DCs and this was intrinsic to the DCs, as it was replicated amongst the progeny of CCR2^−/− BM in mixed WT:CCR2^−/− BM chimeras.

Like CD103⁺ DCs, CD103⁻CD11b⁺ DCs efficiently induced the proliferation of antigen-specific CD4⁺ T cells after antigen loading ex vivo. Significantly, the CCR2-expressing CD103⁻CD11b⁺ DCs induced the polarisation of T_h17 cells in vitro, a function previously associated with CD103⁺CD11b⁺ DCs. This idea was based on evidence from mice lacking IRF4, Notch-2 or human langerin in CD11c⁺ cells, which showed concomitant, selective defects in CD103⁺CD11b⁺ DCs and T_h17 cells in the intestine ^22,36,46,47. Importantly our novel population of T₁₇ inducing, CCR2⁺CD103⁻CD11b⁺ DCs expressed IRF4 and their numbers were reduced in CD11c-cre-IRF4^fl mice, suggesting that this previously unrecognised subset may contribute to the impairment of T_h17 cell generation in these animals. Indeed a very recent study has demonstrated that the role of CD103⁺CD11b⁺ DCs in T_h17 cell homeostasis in the LP may be to act as an accessory cell, rather than to interact with T cells in a cognate manner⁴⁷. More work is clearly warranted to explore how these various subsets of DCs may co-operate in the priming of T cell responses in vivo. The presence of apparently intrinsically pro-inflammatory cells amongst the CD103⁻ DC population could also help explain why CD103⁻ “DCs” were often previously associated with inflammation when less well-defined cell populations were used ^43,48. Finally it should be noted that CD103⁻CD11b⁺ DCs have been shown to drive T_h17 polarisation in other tissues such as the lung ²². As the expression of CD103 by CD11b⁺ DCs is unique to the intestine, it remains possible that the T_h17 polarising CCR2⁺CD103⁻CD11b⁺ DCs we have identified are intermediate stages within the development of CD103⁺CD11b⁺ DCs, whose ultimate differentiation involves the loss of CCR2 and acquisition of CD103, perhaps following their conditioning by the intestinal microenvironment ⁴⁹.

While previous studies have correlated the ability of intestinal DCs to prime T_h17 cells with their production of IL-6 ³⁶, we found similar frequencies of IL-6 producing cells amongst CCR2⁻ and CCR2⁺CD103⁻ DCs. However the CCR2-expressing DCs included more cells containing intracellular IL-12/IL-23p40, both in steady state and even more markedly after TLR4 ligation in vitro. As there were no differences in the ability of the various subsets of CD103⁻CD11b⁺ DCs to prime T_h1 responses in vitro, we propose that the increased levels of IL-12/IL-23p40 in the CCR2-expressing CD103⁻CD11b⁺ DCs reflect enhanced production of IL-23 and that this explains their T_h17 polarising properties. However the difficulties associated with obtaining high numbers of DCs after culture in isolation, together with the current lack of a mouse model which lacks CCR2⁺CD103⁻CD11b⁺ DCs specifically, mean that further work is required to confirm this hypothesis directly.

The potential clinical implications of our results are highlighted by our finding of similar heterogeneity of CCR2 expression amongst CD103⁻ SIRPα⁺ DCs in steady state human intestine, a population that appears to be equivalent to CD103⁻CD11b⁺ DCs in mice ²². It will be of interest to examine the function of this subset particularly in disorders associated with T_h17 cells such as Crohn’s disease. Furthermore, our results showing that CCR2 dependence is not a reliable indicator of monocytic origin and that DC and mϕ are truly derived from distinct precursors raise important issues that need to be considered in the clinical situations where monocyte derived cells and myeloid cell growth factors are being trialled.

In conclusion, we show definitively that CD103⁻CD11b⁺ MPs in the murine intestinal LP include a population of genuine DCs which are derived exclusively from DC committed precursors, rather than Ly6C^hi monocytes as previously assumed. This subset of DCs is itself heterogeneous, containing a population of CCR2-expressing DCs with pro-inflammatory properties. These results provide novel insights which may help explain recent discrepancies on the origins and nature of CD103⁻CD11b⁺ MPs. An analogous subset is present in human intestine, indicating that this heterogeneity of DCs may be relevant across species barriers.

Methods

Mice and Human Tissues

Wild type (WT) C57Bl/6 (B6) mice were purchased from Harlan Olac (Bicester, UK). CX3CR1^+/gfp mice ⁵⁰ and C57Bl/6.SJL (CD45.1⁺) mice were bred originally at Lund University. CCR2^−/− mice ⁵¹ mice were obtained from Professor R. Nibbs (University of Glasgow, UK). OTII mice were bred in house. All these strains were maintained at the University of Glasgow animal facilities. Flt3L^{−/− 52}, CD11c-cre × IRF4^floxed mice ³⁶, and C57Bl/6 WT mice were bred and maintained at the VIB Ghent University, Ghent. CD11c-cre × IRF4^floxed mice were also bred and maintained at Lund University. All mice were backcrossed for at least 9 generations on to the B6 background, maintained under specific pathogen free (SPF) conditions and were used between 6 and 12 weeks of age. Animal experiments were performed in accordance with UK Home Office guidelines.

Histologically normal samples of human colon were obtained from patients undergoing surgical resections for adenocarcinoma. Tissues were obtained with informed consent and experiments performed in accordance with the Biorepository NHS GGC ethics application number 65.

Murine cell isolation

Lamina propria cells were obtained from murine intestines by enzymatic digestion as described previously ^5,28. Cells were isolated from mesenteric lymph nodes by enzymatic digestion with 1mg/ml collagenase D (Roche) in calcium magnesium free (CMF) Hank’s balanced salt solution (HBSS; Gibco, Invitrogen) for 45 minutes. After isolation, cells were passed through a 100μm and a 40μm filter before use (Corning).

Human cell isolation

Underlying fat and muscle layers were removed, tissue was washed in CMF HBSS 2% FCS (Sigma) and cut into 0.5 cm sections. The tissue was then shaken vigorously in 10ml HBSS/2% FCS and the supernatant discarded. To remove the epithelial layer, 10 ml fresh CMF HBSS containing 2mM EDTA (Sigma) was added, the tube placed in a shaking incubator for 15mins at 37°C, before being shaken vigorously and the supernatant discarded. The intestinal tissue was washed in CMF HBSS and the EDTA step repeated twice more. The remaining tissue was digested with pre-warmed complete RPMI 1640 supplemented with 2 mM L-glutamine, 100μg/ml penicillin, 100μg/ml streptomycin, 1.25μg/ml Fungizone (all Gibco, Invitrogen), and 10% FCS containing collagenase VIII (1mg/ml, Sigma), Collagenase D (1.25mg/ml, Roche), Dispase (1mg/ml, Gibco, Invitrogen) and DNase (30μg/ml, Roche) for 45 minutes in a shaking incubator at 37°C. The resulting cell suspension was removed and the digestion was repeated until all tissue was dispersed. Cell suspensions were passed through a 40μm cell strainer (BD Biosciences), washed twice in complete RPMI and kept on ice until use.

Flow cytometric analysis and sorting

Cells were stained at 4°C in the dark as described previously ²⁸ using the antibodies listed in supplementary experimental procedures online. For intracellular cytokine staining, whole LP digests were incubated for 4.5 hours with 5μg/ml brefeldin A and 2μM monensin (both Biolegend) in the presence or absence of 100ng LPS from Salmonella typhimurium (Sigma) before fixation and permeabilisation. In all analyses, following doublet exclusion, live cells were identified using 7-AAD (Biolegend) or fixable viability dye (eBioscience). Data were acquired on an LSR II, Fortessa, FACSAria I or FACSAria III (BD Biosciences) and analysed using FlowJo software (Tree Star Inc.).

Adoptive transfer of bone marrow precursors

For pre-DCs, WT mice were injected with 2×10⁶ flt3L secreting B16 tumour cells subcutaneously (a kind gift from Dr Oliver Pabst, Hannover, Germany) and 10-14 days later BM was isolated and RBC lysed (Stem Cell Technologies). Cells were labelled with eFluor450 CellTrace Violet proliferation dye (eBioscience) and pre-DCs identified as Lin⁻CD11c^intCCR9⁻B220^loCD135⁺ cells. 7×10⁵ FACS sorted pre-DCs were injected into unmanipulated congenic recipients, or into mice that had received 1.2% DSS ad libitum in the drinking water for 3 days prior to transfer. For monocyte transfers, 1×10⁶ FACS-purified Ly6C^hi monocytes (CD11b⁺CD117⁻Ly6G⁻Ly6C^hiCX3CR1^int) from the BM of CX3CR1^+/gfpCD45.1⁺/CD45.2⁺ mice were transferred into CD45.2 CCR2^−/− mice.

Antigen specific T cell proliferation in vitro

DC subsets were FACS-purified and pulsed with 2mg/ml ovalbumin (Grade VI, Sigma) for 2 hours before being washed extensively and co-cultured for 4 days with CFSE-labelled, naïve CD62L^hiCD25⁻CD4⁺ T cells FACS-purified from the lymph nodes of OTII mice. For intracellular cytokine staining, cultured cells were placed in a stimulation cocktail (eBioscience) containing PMA, ionomycin, brefeldin A and monensin for 4.5 hours, before fixation, permeabilisation and staining with antibodies against IL-17a, IFNγ and FoxP3 .

Quantitation of gene expression by real time reverse transcription PCR

Total RNA was purified from sorted LP cells using the RNeasy Micro kit (Qiagen). RNA was reverse transcribed to cDNA and gene expression was assayed by quantitative reverse transcription PCR (qRT-PCR) as described previously ¹⁶. The primer sequences used are detailed in Supplementary experimental procedures online.

Generation of WT:CCR2^−/− mixed BM chimeras

CD45.1/CD45.2 WT mice received two doses of 5Gy, 2 hours apart before receiving 5×10⁶ BM cells i.v. from CD45.1 WT and CD45.2 CCR2^−/− BM at a 50:50 ratio. Chimerism was assessed 8 weeks after reconstitution.

Assessment of phagocytosis

3×10⁶ cells were assessed for phagocytosis of pHrodo E. coli bioparticles (Molecular Probes, Life Technologies) according to the manufacturer’s guidelines and analysed by flow cytometry.

Assessment of BrdU incorporation in vivo

Mice were injected once i.p. with 1mg BrdU (BD Biosciences) and in some experiments received 0.8mg/ml BrdU (Sigma) in their drinking water for 3 days. The incorporation of BrdU by isolated cells was assessed using the BD BrdU Flow kit (BD Biosciences).

Statistical Analysis

Results are presented as means ±1 standard deviation unless otherwise stated and groups were compared using a Student’s t test, or for multiple groups, a one-way ANOVA followed by a Bonferroni post test using Prism Software (GraphPad Software, Inc.).

Supplementary Material

Supplementary Figures 1 and 2

Supplementary Figure 1: Phenotypic characterization of mononuclear phagocytes in colonic LP and MLNs.

(a) Live leukocytes (7AAD⁻CD45⁺) were identified in colonic LP digests and analysed for expression of CD11c and MHCII. Macrophages were excluded based on F4/80 expression and the resulting CD11c⁺F4/80⁻ MPs then analysed for CD103 and CD11b expression. Data are representative of at least 5 experiments (b) DC populations were identified in the LP of the duodenum, jejunum ileum and colon and the results shown are the frequencies of each DC subset as a % of total live CD45⁺CD11c⁺MHCII⁺ cells. (c) Frequencies of CD64⁺ mϕs as a percentage of live CD45⁺CD11b⁺ cells in the duodenum, jejunum, ileum and colon. Data are from a single experiment with n=4 *p<0.05, **p<0.01, ***p<0.005, vs the equivalent population in duodenum. One way ANOVA with Bonferroni post-test. (c) Live CD45⁺B220⁻ cells were identified in MLN digests and analysed for expression of CD11c and MHCII. MLN MPs were identified as migratory (mig) (CD11c⁺MHCII^hi) or resident (res) (CD11c⁺MHCII^int) and after exclusion of CD64⁺ mϕ, mig and res DCs were analysed for expression of CD103 and CD11b. (d) Proportions of each CD64⁻ mig. DC population as a percentage of total mig. DCs, with each point representing a mouse and the horizontal line indicating the mean. Data are representative of at least 10 independent experiments.

Supplementary Figure 2: Identification and FACS-purification of BM precursors for adoptive transfer.

(a) Ly6C^hi monocytes were identified amongst BM leukocytes of CX3CR1^+/GFP mice as CD11b⁺ Ly6G⁻Ly6C^hiCD117⁻CX3CR1^int. FACS-purified Ly6C^hi monocytes used in adoptive transfer experiments were uniformly CD11b⁺ Ly6G⁻Ly6C^hiCD117⁻. Numbers represent purity of Ly6C^hi monocytes as a percentage of total sorted cells. Data are representative of at least 2 independent experiments. (b) Pre-DCs were identified amongst single BM leukocytes as lineage⁻ (CD3, CD19, CD49b, MHCII, and CD11b), CD11c⁺, B220⁻ and CCR9⁻. FACS-purified pre-DCs used in adoptive transfer experiments were uniformly Lin⁻CD11c⁺CCR9⁻B220⁻. Numbers represent purity of pre-DCs as a percentage of total sorted cells. Data are representative of at least 10 independent experiments. (c) Pre-DCs (black line) were homogeneously CD135⁺, CD117⁻ and CD172α^int compared with isotype control or lineage⁻CD45⁺ cells (CD117 control only) (shaded grey). (d) Expression of MHCII on pre-DCs before transfer (black line) and on total live CD45⁺ cells in the SI LP. (e) 7×10⁵ CD45.2⁺Violet⁺ pre-DCs were transferred into resting WT CD45.1⁺ mice and 1, 3, 5 or 7 days later, the SI LP of recipient mice was examined for the presence of donor-derived cells. Representative contour plots of CD11c and MHCII expression on live-gated host cells or donor-derived cells, 1, 3, 5 and 7 days after transfer. Numbers indicate the frequencies of CD11c⁺MHCII⁺ cells as a percentage of total donor-derived cells. Data are representative of either a single experiment (1 & 3 days) or 3 independent experiments (5 & 7 days) with 2-4 recipient mice per time point per experiment.