Exclusive CX3CR1 dependence of kidney DCs impacts glomerulonephritis progression (original) (raw)
Dependence of kidney DCs and MPs on CX3CR1 in healthy mice. CX3CR1 is abundantly expressed on most kidney APCs, whereas other immune cells constituted only minor fractions among CX3CR1+ kidney cells (ref. 8 and Supplemental Figure 1, A–C; supplemental material available online with this article; doi:10.1172/JCI70143DS1). The functional role of CX3CR1 in kidney APCs is unclear. When we compared kidney sections of CX3CR1GFP/+ mice (expressing a GFP reporter in CX3CR1+ cells) and of CX3CR1GFP/GFP mice (expressing the GFP reporter and deficient for CX3CR1) by fluorescence microscopy, an obvious reduction of GFP+ cells in the absence of CX3CR1 was noted (Figure 1A). Analysis of kidney single cell suspensions by flow cytometry (gating strategy shown in Supplemental Figure 1D) revealed that exclusively the CX3CR1+ APCs expressing CD11c, which have been shown to possess DC functionality (6), were reduced by 75% in CX3CR1GFP/GFP mice, whereas CD11c– MPs were slightly, albeit not statistically significantly, enhanced (Figure 1B). Total DC and MP numbers did not differ between WT and CX3CR1GFP/+ mice (Supplemental Figure 2A), indicating that the latter could serve as CX3CR1-competent controls.
CX3CR1 specifically regulates the abundance of DCs in the kidney. (A) Representative kidney sections of CX3CR1GFP/+ reporter mice and CX3CR1GFP/GFP-deficient mice (original magnification, ×200). Arrows point to GFP+ cells. (B) CD11c versus MHC-II staining of CD45+GFP+ cells from CX3CR1GFP/+ reporter mice (left) and quantification of DCs (MHC-II+CD11c+) and MPs (MHC-II+CD11c–) (right) in CX3CR1GFP/+ (GFP/+) and CX3CR1GFP/GFP mice (GFP/GFP). Results of 5 individual experiments were combined. (C) Absolute numbers of GFP+ DCs in various organs and tissues of CX3CR1GFP/+ and CX3CR1GFP/GFP mice, given as total DCs per organ. Results are representative of 2 individual experiments, with 3 to 4 mice per group. ingLN, inguinal LN. (D) Relative CX3CL1 expression in different organs compared with expression levels in the kidneys. Statistical significance was tested with (B) unpaired Student’s t test or (C and D) 1-way ANOVA with Bonferroni multiple-comparison test. **P < 0.01, ***P < 0.001.
To study whether the reduction in DC numbers was unique to the kidney, we compared their abundance in different tissues from CX3CR1GFP/+ and CX3CR1GFP/GFP mice. There was no reduction in liver, lung, heart, bladder, lymph nodes, or spleen (Figure 1C). DCs were less frequent in CX3CR1-deficient mice in the small intestine, but their absolute numbers were more than an order of magnitude lower than in 1 kidney (Figure 1C). The spleens of CX3CR1GFP/GFP mice contained approximately 40% more GFP+ DCs than those of CX3CR1GFP/+ controls (Figure 1C), which might be explained by different homing of DC precursors.
CD11c– GFP+ MPs were substantially reduced in the intestines of CX3CR1-deficient mice (Supplemental Figure 2B), consistent with a recent report (22). There was no reduction of CD11c– GFP+ cells in kidney, liver, lung, heart, bladder, dermis, epidermis, bone marrow, spleen, and brain (Supplemental Figure 2C).
We speculated that the particularly strong CX3CR1 dependence of DCs in the kidney might be due to high expression of its ligand, CX3CL1, in this organ. Indeed, when we determined CX3CL1 by quantitative RT-PCR, its expression in the kidney was much higher than that in other organs except the small intestine (Figure 1D), providing a plausible explanation as to why DCs depended on CX3CR1 only in these organs (Figure 1C). There were no differences in CX3CL1 expression between CX3CR1-deficient and competent mice (Supplemental Figure 2D), ruling out feedback regulation of ligand expression by the receptor.
Dependence of kidney DCs and MPs on CX3CR1 in NTN. To study whether kidney DCs depended on CX3CR1 also under inflammatory conditions, we induced NTN, a widely studied model for crescentic GN. Ten days after disease induction, GFP+ DCs were scarcer in the kidneys of CX3CR1GFP/GFP mice compared with CX3CR1GFP/+ mice (Figure 2A). Of note, there was also a slight, albeit not statistically significant, reduction of renal GFP+ MP numbers at that time point (Figure 2A). The small numbers of CX3CR1+ non-APCs, such as T cells, NK cells, and mast cells, differed very little between CX3CR1GFP/+ and CX3CR1GFP/GFP mice, both under homeostatic conditions and in NTN (Supplemental Figure 1, A and B).
Dependence of kidney DCs and MPs on CX3CR1 in NTN. (A) Absolute numbers of GFP+ DCs and MPs per kidney in CX3CR1GFP/+ and CX3CR1GFP/GFP mice on day 10 after NTN induction. Results are combined from 4 individual experiments. (B) Ly6C expression on GFP+ bone marrow cells from CX3CR1GFP/+ and CX3CR1GFP/GFP mice before transfer into nephritic WT mice. (C) Representative FACS plots of CD11c and MHC-II expression on GFP+ cells recovered from kidneys of nephritic WT mice, which had received 106 GFP+ bone marrow cells from CX3CR1GFP/+ or CX3CR1GFP/GFP mice 96 hours before. (D) Proportion of GFP+ cells among CD45+ cells in the kidneys of nephritic WT mice 48 and 96 hours after transfer of 106 GFP+ bone marrow cells from CX3CR1GFP/+ or CX3CR1GFP/GFP mice. (E) Proportion of GFP+ cells among CD45+ cells in the spleens of nephritic WT mice 96 hours after transfer of 106 GFP+ bone marrow cells from CX3CR1GFP/+ or CX3CR1GFP/GFP mice. Data points represent (A and E) individual mice or (D) organs. Results are representative 2 individual experiments, with 3 to 4 mice per group. Statistical significance was tested with (A) the unpaired Student’s t test or (D) the 2-tailed Mann-Whitney test.
We next asked whether the reduction of kidney DCs in nephritic CX3CR1GFP/GFP mice was only a consequence of the reduced DC numbers under homeostatic conditions (Figure 1B) or whether the recruitment of DC precursors under inflammatory conditions also depended on CX3CR1. We addressed this question by adoptively transferring 106 GFP+ bone marrow cells from CX3CR1GFP/+ and CX3CR1GFP/GFP mice into WT mice 1 day after NTN induction. More than 60% of the GFP+ bone marrow cells expressed Ly6C, which is characteristic of inflammatory monocytes (Figure 2B). Influx of GFP+ cells into the kidneys was analyzed 2 and 4 days after transfer. GFP+ cells expressing the DC markers CD11c and MHC-II were detected in inflamed kidneys (Figure 2C). Importantly, far fewer CX3CR1-deficient cells entered nephritic kidneys at both time points (Figure 2D), whereas more of them were recovered from the spleens (Figure 2E). This indicates that CX3CR1 contributes to entry of DC precursors into the inflamed kidney. In the absence of CX3CR1, these cells relocated to the spleen, similar to our observations in healthy mice (Figure 1C). We did not detect the marker Ly6C on immigrated DCs (data not shown). This may be due to the rapid loss of this marker soon after tissue entry of Ly6C+ monocytes, as recently shown in the lung (38).
CX3CR1 deficiency attenuates NTN. We recently showed that kidney DC maturation stimulates the intrarenal DTH that drives NTN (13). The dearth of DCs in nephritic CX3CR1GFP/GFP mice suggested that DTH should be reduced in these mice. Indeed, flow cytometric analysis of kidney single cell suspensions (gating strategies shown in Supplemental Figure 1, D–F) on day 10 after disease induction revealed reduced numbers of IFN-γ+ CD4+ T cells (Figure 3A) and of TNF-producing DCs and MPs in CX3CR1-deficient mice (Figure 3B), suggesting an attenuated intrarenal DTH response. IL-17+ CD4+ T cells were only slightly and not significantly reduced (Supplemental Figure 3B), consistent with the recent finding that Th17 cells are more important in the early disease phase (14). Other early immune mediators were also not affected by CX3CR1 deficiency, as evidenced by unchanged IFN-γ, TNF, CXCL10, and CXCL16 expression in CX3CR1GFP/+ and CX3CR1GFP/GFP mice after 24 hours (Supplemental Figure 3A).
CX3CR1 deficiency reduces intrarenal Th1 response and improves kidney function. (A) Absolute numbers of IFN-γ–producing CD4+ T cells and (B) of TNF-producing DCs and MPs per kidney of CX3CR1GFP/+ reporter mice and CX3CR1GFP/GFP-deficient mice 10 days after NTN induction, as determined by intracellular flow cytometry. In A, cell numbers in 4 individual experiments were given as percentage of the mean cell number in the CX3CR1-competent groups. (C) Creatinine clearance in CX3CR1GFP/+ and CX3CR1GFP/GFP mice 10 days after NTN induction. (D) Blood urea nitrogen in CX3CR1+/+ and CX3CR1GFP/GFP mice 10 days after NTN induction. (E) Albumin excretion normalized to creatinine by CX3CR1GFP/+ and CX3CR1GFP/GFP mice 10 days after NTN induction. Urine was collected for 12 hours. Data points represent individual mice, and results (A–C) are combined from 2 to 4 individual experiments, with 2 to 5 mice per group, or (D and E) are representative of 2 individual experiments, with 10 mice per group. Statistical significance was tested with the 2-tailed Mann-Whitney test.
Functional parameters like creatinine clearance, blood urea nitrogen, and proteinuria were improved in CX3CR1GFP/GFP mice (Figure 3, C–E), and histological examination of kidney sections revealed fewer tubular casts and crescents in CX3CR1GFP/GFP mice on day 10 (Supplemental Figure 3C), indicating that the loss of CX3CR1 attenuates NTN. On day 15, the histological features of kidney damage were even more pronounced (Figure 4, A–C). CX3CR1GFP/GFP mice also featured fewer periglomerular F4/80+ cells (Figure 4, D and E), indicating a reduction in DCs and/or MPs. Despite their reduced numbers (Figure 2A), kidney DCs of CX3CR1GFP/GFP mice still formed periglomerular infiltrates (6) in NTN (Figure 4F), indicating that CX3CR1 was dispensable for intrarenal APC migration toward inflamed glomeruli.
CX3CR1 deficiency attenuates NTN. (A) Kidney sections of CX3CR1+/+ and CX3CR1GFP/GFP animals 15 days after NTS challenge (Masson trichrome; original magnification, ×200). (B) Percentage of crescentic glomeruli in histological sections of CX3CR1+/+ and CX3CR1GFP/GFP mice on day 15 after disease induction. (C) Tubular casts per high-power field (hpf) (original magnification, ×100) in CX3CR1+/+ and CX3CR1GFP/GFP mice 15 days after NTS challenge. (D) Representative microphotographs of immunostained F4/80+ cells in kidney cortices from CX3CR1+/+ and CX3CR1GFP/GFP mice on day 15 after NTN induction (original magnification, ×200). (E) Quantification of infiltrating F4/80+ cells in CX3CR1+/+ and CX3CR1GFP/GFP animals 15 days after NTS challenge. (F) Immunofluorescence microscopy of kidney sections from nephritic CX3CR1GFP/+ and CX3CR1GFP/GFP mice on day 10 after NTN induction (red [autofluorescence], green [GFP]). Scale bar: 50 μm. Data points represent individual mice, and (D–F) results are representative of 2 individual experiments, with 12 to 16 mice per group. Statistical significance was tested by 2-tailed Mann-Whitney test.
Cell-intrinsic DC functions are independent of CX3CR1 expression. To understand how CX3CR1 deficiency attenuated intrarenal DTH and nephritis symptoms, we asked whether DCs were functionally defective in the absence of this receptor. We first examined the systemic Th1 response against sheep Ig, which is induced by DCs in lymphatic tissues after injection of the nephrotoxic sheep serum (39). This response was unaltered, as evidenced by similar subcutaneous DTH response against sheep Ig (Figure 5A) and by similar production of the Th1 and Th17 cytokines, IFN-γ, TNF, and IL-17, by splenocytes (Figure 5B). These findings excluded the possibility that the attenuated intrarenal DTH response in CX3CR1GFP/GFP mice resulted from a failure to activate a nephritogenic T cell response.
Cell-intrinsic DC functions are unaltered by CX3CR1 deficiency. (A) DTH reaction in the skin of CX3CR1GFP/+ reporter mice and CX3CR1GFP/GFP-deficient mice, rechallenged by footpad injection with NTS. Depicted is footpad swelling in comparison with the contralateral footpad injected with vehicle control. (B) IFN-γ, IL-17, and TNF concentration in the supernatant of 24-hour splenocyte culture from nephritic CX3CR1GFP/+ and CX3CR1GFP/GFP mice rechallenged with NTS. 107 total splenocytes were cultured for 24 hours in 1 ml medium and 1:200 NTS. (C) MFI of CD86 and MHC-II expression on kidney DCs from CX3CR1GFP/+ mice and CX3CR1GFP/GFP mice, determined by flow cytometry. (D) Representative FACS plots of the in vivo uptake of fluorescently labeled OVA by kidney DCs of nephritic CX3CR1GFP/+ and CX3CR1GFP/GFP mice gated on CD45+MHC-II+CD11c+ cells. Numbers show percentages of OVA+ DCs. (E) Representative histograms of T cell proliferation and CFSE dilution and division indices (DI) in a coculture of OVA-specific T cells and kidney DCs from nephritic CX3CR1GFP/+ and CX3CR1GFP/GFP mice injected with 1 mg OVA 1 hour prior to cull. (F) IFN-γ and IL-17 concentrations in supernatants of 72-hour cocultures from E. Data points represent individual mice, and results are representative of 2 to 4 experiments, with 3 to 4 mice per group. Statistical significance was tested with the unpaired Student’s t test.
We therefore focused on intrarenal DCs. CX3CR1-deficient and -competent kidney DCs from nephritic mice expressed equal levels of costimulatory molecules and MHC-II (Figure 5C). Next, we studied their ability to capture antigen by injecting the model antigen, OVA, a 45-kDa protein that can constitutively pass the glomerular filter (40). Regardless of CX3CR1 expression, DCs took up similar amounts of OVA (Figure 5D). To examine the principal DC function, the activation of naive T cells, we injected CX3CR1-deficient and CX3CR1-competent nephritic mice with OVA and cocultured kidney DCs with OVA-specific Th cells (OT-II cells). CX3CR1GFP/+ and CX3CR1GFP/GFP DCs elicited similar proliferation profiles and cytokine production by OT-II cells (Figure 5, E and F). Therefore, the observed differences in disease severity were not due to DC-intrinsic functional defects, supporting their reduced numbers as an underlying reason.
DCs in the renal cortex are especially dependent on CX3CR1. When we reexamined kidney sections of healthy CX3CR1GFP/+ and CX3CR1GFP/GFP mice by immunofluorescence microscopy, the paucity of GFP+ cells in the latter mice seemed to preferentially affect the kidney cortex (Figure 6A). To verify this histological impression in a quantitative manner and to distinguish between DCs and MPs, we surgically separated the renal medulla from the cortex, produced single cell suspensions by separate collagen digestion, and analyzed these by flow cytometry. The cortex of CX3CR1GFP/+ mice contained 3 times more GFP+ cells than the medulla, and 80% of them expressed the DC marker CD11c, whereas less than 40% of GFP+ cells in the medulla did so (Figure 6, B and C). Consequently, the absolute numbers of GFP+ DCs were about 6 times higher in the cortex than in the medulla (Figure 6D). In CX3CR1GFP/GFP mice, DC numbers were reduced 9 fold from 200 × 103 to 22 × 103 cells per kidney cortex, whereas medullary DCs were reduced 3.5 fold from 38 × 103 to 11 × 103 cells per kidney (Figure 6D). MP numbers were increased 2.5 fold in the cortices of CX3CR1GFP/GFP mice (Figure 6E). Of note, the mice used in this experiment were younger than those used in the other experiments (Figure 1 and Figure 6, G and H), explaining the relatively low absolute DC numbers. Nevertheless, DC numbers were similarly reduced, indicating that the kidney DC deficiency in CX3CR1-deficient mice was age independent.
DCs in the renal cortex particularly depend on CX3CR1. (A) Histological quantification of GFP+ cells in cortical and medullary kidney sections of healthy CX3CR1GFP/+ reporter mice and CX3CR1GFP/GFP-deficient mice. Each data point represents 1 high-power field (original magnification, ×200). Statistical significance was calculated by Mann Whitney test. (B) Absolute numbers of GFP+ cells in the cortices and medullas of 1 kidney of healthy CX3CR1GFP/+ mice (gray bars) and CX3CR1GFP/GFP mice (black bars), determined by flow cytometry. (C) Representative FACS plots of CD45+GFP+ cells in kidney cortices and medullas of CX3CR1GFP/+ mice. (D and E) Absolute numbers of (D) DCs and (E) MPs in the renal cortices and medullas of healthy CX3CR1GFP/+ mice (gray bars) and CX3CR1GFP/GFP mice (black bars). (F and G) Absolute numbers of (F) DCs and (G) MPs in the cortices and the medullas of 1 kidney of nephritic CX3CR1GFP/+ mice (gray bars) and CX3CR1GFP/GFP mice (black bars) on day 10 after NTN induction. Results are representative of 2 to 3 individual experiments, with 3 to 4 mice per group. Statistical analysis was performed using 1-way ANOVA with Bonferroni multiple-comparison. **P < 0.01, ***P < 0.001.
The kidney cortices of nephritic CX3CR1GFP/+ mice contained 416 × 103 GFP+ DCs, compared with 164 × 103 DCs in CX3CR1GFP/GFP mice (60% reduction), whereas only 96 × 103 DCs were seen in the medullas of CX3CR1-competent mice and 44 × 103 DCs were seen in those of CX3CR1-deficient mice (55% reduction) (Figure 6F). Thus, even though proportional DC reduction was similar, the absolute deficit of kidney DCs in nephritis was more prominent in the cortex (–252 × 103 DCs) than in the medulla (–52 × 103 DCs). Furthermore, there were approximately 32 × 103 fewer MPs in the cortices and approximately 20 × 103 fewer medullary MPs in CX3CR1-deficient nephritic mice (Figure 6G). Taken together, our findings demonstrated that, under homeostatic conditions, CX3CR1 primarily affected DCs in the cortex, and, in the condition of nephritis, affected both cortical and medullary DCs and to a lesser extend also MPs.
Cortical, but not medullary, DCs can stimulate Th cells in GN. We reasoned that the proximity of cortical DCs to the glomeruli might render these cells particularly relevant in GN. When we compared the phenotype of medullary and cortical APCs in healthy mice, we observed similar expression of CD11b, F4/80, and CX3CR1 and higher CD11c expression on cortical CX3CR1+ cells (Figure 7A). For the most part, cortical and medullary DCs expressed similar levels of MHC-II and of the costimulatory molecules CD80, CD86, and CD40 under homeostatic conditions (Figure 7B), notwithstanding some variability of expression levels between experiments. In nephritic mice, expression of CD40 and MHC-II on cortical DCs slightly exceeded that on medullary DCs in 3 out of 5 experiments, consistent with higher maturation of the former (Figure 7B).
Cortical, but not medullary, DCs can stimulate Th cells in GN. (A) Flow cytometric analysis of CX3CR1+ APCs from kidney cortices (gray lines) and medullas (black dashed lines) of CX3CR1GFP/+ reporter mice for expression levels of CD11c, CD11b, F4/80, CX3CR1, and CD103. (B) MFI, reflecting expression levels of MHC-II, CD40, CD86, and CD80 on cortical (white bars) and medullary (black bars) DCs in healthy mice (day 0) or mice on day 7 or 10 after NTN induction. (C) Representative CFSE dilution profiles, reflecting T cell proliferation in a 72-hour coculture of OVA-specific CD4+ T cells and OVA-loaded cortical, medullary, or splenic DCs derived from healthy (gray solid) or 7-day nephritic (black line) WT mice. 2 × 104 DCs were loaded with 1 mg/ml OVA for 2 hours, washed, and cocultured with 1.5 × 105 OT-II cells for 72 hours. (D) Division indices of OT-II cells in coculture from C. (E) IFN-γ and IL-17 concentrations in 200 μl medium after 72 hours coculture, as in C. Results are representative of at least 2 individual experiments, with at least 3 mice per group. Statistical significance was analyzed applying the paired Student’s t test. **P < 0.01.
NTN is driven by a Th cell–dependent intrarenal DTH (11). Therefore, we cocultured cortical and medullary DCs with the model antigen, OVA, and OT-II cells. OVA-loaded cortical and medullary DCs from healthy mice caused little T cell proliferation compared with control splenic DCs (Figure 7C). By contrast, OVA-loaded DCs from the cortices of nephritic mice induced strong OT-II cell proliferation (Figure 7, C and D) and secretion of IFN-γ and IL-17 (Figure 7E). Remarkably, medullary DCs from nephritic kidneys still only elicited minimal OT-II cell proliferation and cytokine production (Figure 7, C–E). These results suggested that Th cell stimulation during NTN depends mainly on cortical DCs, implying that only these DCs drive NTN progression.
Medullary DCs poorly process antigen for MHC-II presentation. To understand why medullary DCs poorly stimulated Th cells, we first examined their survival in vitro. A similar fraction of cortical and medullary DCs survived a culture period of 24 hours (cortical DCs: 55.4% ± 0.7% survival; medullary DCs: 58.5% ± 2.4% survival), excluding selective death as underlying reason. DC maturation parameters differed little (Figure 7B). We therefore speculated about distinct antigen handling as an underlying reason and first asked whether medullary DCs were unable to endocytose antigen. However, medullary DCs captured slightly more fluorescent OVA than cortical DCs in an in vitro culture both under homeostatic conditions (Figure 8, A and B) and in NTN (Figure 8B). To test whether medullary DCs might not have antigen access in vivo, we injected fluorescent OVA i.v. into mice and determined its uptake by medullary and cortical DCs. More medullary than cortical DCs captured OVA under homeostatic conditions, and they also took up more filtered antigen per cell (Figure 8, C–E). On day 10 of nephritis, when the mice are heavily proteinuric, the proportion of antigen-containing cortical DCs was as high as that in medullary DCs (Figure 8, C–E). However, the antigen uptake per cell was 3-fold higher in medullary DCs than in cortical DCs (Figure 8, D and E). These findings exclude the possibility that impaired antigen uptake explains the functional defect of medullary DCs.
Medullary DCs do not process antigen for MHC-II presentation. (A) Uptake of fluorescently labeled OVA by sorted CX3CR1+ APCs from kidney cortices (white squares) and medullas (black squares) of healthy CX3CR1GFP/+ reporter mice. Sorted GFP+ cells were incubated with OVA for 20 minutes at 37°C, washed, and analyzed by flow cytometry. (B) Sorted cortical (white bars) and medullary (black bars) DCs from healthy or 10-day nephritic mice were incubated with 10 μg/ml OVA, as in A. MFI of OVA+ cells, reflecting average amount of antigen uptake per cell. (C) Percentage of OVA+ DCs in cortex (white bars) and medulla (black bars) 30 minutes after i.v. injection of fluorescently labeled OVA into healthy or 10-day nephritic mice. (D) Representative FACS plots of the in vivo OVA uptake of cortical and medullary CD45+MHC-II+ cells in healthy or 10-day nephritic mice, 30 minutes after i.v. injection of Alexa Fluor 647–conjugated OVA. (E) MFI of OVA+ cells in cortex and medulla 30 minutes upon i.v. injection of OVA, as in C. (F) MFI of cortical and medullary DCs, indicating cleaved DQ-OVA after 20 minutes of incubation with 200 μg/ml DQ-OVA. (G) Analysis of cleaved DQ-OVA per endocytosed Alexa Fluor 647–conjugated OVA in cortical and medullary DCs, after 3 hours of incubation with 25 μg/ml DQ-OVA and Alexa Fluor 647–conjugated OVA. (H) mRNA for Cathepsin H, invariant chain, and H2-DM in cortical and medullary DCs from 7-day nephritic mice, relative to GAPDH expression. Results represent 2–3 experiments, with 3 to 5 mice per group. Statistical analysis (B–E) by 1-way ANOVA with Bonferroni test or (F and H) by paired Student’s t test. *P < 0.05, **P < 0.01, ***P < 0.001.
We next speculated that medullary DCs might function like MPs, which capture more antigen than DCs but degrade it before it can be presented to T cells (41). To test this, we offered the DCs DQ-OVA, which becomes fluorescent once it is cleaved. Medullary DCs cleaved somewhat more DQ-OVA than cortical DCs (Figure 8F), but this increase correlated with their higher endocytotic activity (Figure 8G). Thus, medullary DCs did not possess the high degradative activity of MPs. This left the explanation that the antigen processing and MHC-II loading machinery is defective in medullary DCs. While the antigen-degrading enzyme cathepsin H was not differentially expressed, there was indeed less expression of invariant chain and H2-DM in medullary DCs, suggesting that medullary DCs are less well equipped to load processed antigen onto MHC-II for presentation to Th cells (Figure 8H).
Medullary DCs regulate innate immunity in the kidney. The inability of medullary DCs to activate Th cells raised the question of what physiological role these cells might play. We reasoned that their location might be particularly suitable for the sentinel function we recently described for kidney DCs in PN (4), because uropathogenic E. coli ascend through the collecting ducts in the medulla. To test this hypothesis, we determined DC production of the chemokine CXCL2, which attracts neutrophils into the infected kidney. Indeed, 3 hours after infection, medullary DCs produced far more CXCL2 than cortical DCs (Figure 9, A–C). MPs, identified by lack of CD11c expression, produced very little CXCL2 (Figure 9A).
Medullary DCs regulate the innate immunity against PN. (A) Representative FACS plots of CXCL2 production by CD45+MHC-II+ cells in kidney cortex and medulla 3 hours after second UPEC instillation. Cells were incubated with brefeldin A for 4 hours at 37°C prior to staining. (B) Percentage of CXCL2-producing cells of CD45+MHC-II+CD11c+ cells in cortex (white squares) and medulla (black squares). (C) MFI for CXCL2 of cortical and medullary CXCL2+ DCs, representing CXCL2 production per cell. (D) CFUs per kidney in CX3CR1GFP/+ reporter mice and CX3CR1GFP/GFP-deficient mice 6 hours after second UPEC instillation. (E) Numbers of neutrophils (PMNs) per kidney in CX3CR1GFP/+ and CX3CR1GFP/GFP mice 6 hours after second UPEC instillation. (F) Representative histogram of CXCL2 staining (black line) of CD45+Ly6G+ neutrophils from CX3CR1GFP/+ and CX3CR1GFP/GFP mice 6 hours after second UPEC instillation, compared with isotype control (gray solid). Cells were incubated with brefeldin A for 4 hours at 37°C prior to staining. Data points represent individual mice; and results are representative of 2 to 3 individual experiments, with 6 mice per group. Statistical significance was analyzed by (B and C) Wilcoxon signed-rank test or (D and E) 2-tailed Mann-Whitney nonparametric test.
We next induced PN in CX3CR1-deficient mice that possess fewer kidney DCs (Figure 1B and Figure 6D) in order to determine consequences for the anti-infectious immune response. Neutrophil numbers in CX3CR1-deficient mice were somewhat reduced at 3 hours after infection (CX3CR1+/+ mice: 81 × 103 ± 17 × 103 neutrophils per kidney; CX3CR1GFP/GFP mice: 54 × 103 ± 17 × 103 neutrophils per kidney; P < 0.01), indicating an initial delay of neutrophil recruitment. However, at 6 hours after infection, bacterial CFUs (Figure 9D) and neutrophil numbers (Figure 9E) did not differ between CX3CR1-deficient and -competent mice. This was not because the fewer DCs in CX3CR1-deficient mice produced more CXCL2 per cell (CX3CR1GFP/+ mice: MFICXCL2 = 7,957 ± 774; CX3CR1GFP/GFP mice: MFICXCL2 = 7,948 ± 487). Instead, we found that neutrophils commenced robust CXCL2 production at 6 hours after infection (Figure 9F). This suggested that the medullary DCs in CX3CR1-deficient mice were sufficient for initiating neutrophil recruitment, and once neutrophils were present in the infected kidney, they produced large amounts of CXCL2. This production compensated for the DC deficiency in the absence of CX3CR1 with respect to antibacterial immune defense.