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Keywords:

  • APCs;
  • DCs;
  • Infectious diseases;
  • Salmonella;
  • T helper cell

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

Control of intracellular Salmonella infection requires Th1 priming and IFN-γ production. Here, we show that efficient Th1 priming after Salmonella infection requires CD11c+CD11bhiF4/80+ monocyte-derived dendritic cells (moDCs). In non-infected spleens, moDCs are absent from T-cell zones (T zones) of secondary lymphoid tissues, but by 24 h post-infection moDCs are readily discernible in these sites. The accumulation of moDCs is more dependent upon bacterial viability than bacterial virulence. Kinetic studies showed that moDCs were necessary to prime but not sustain Th1 responses, while ex vivo studies showed that antigen-experienced moDCs were sufficient to induce T-cell proliferation and IFN-γ production via a TNF-α-dependent mechanism. Importantly, moDCs and cDCs when co-cultured induced superior Th1 differentiation than either subset alone, and this activity was independent of TNF-α. Thus, optimal Th1 development to Salmonella requires the rapid accumulation of moDCs within T zones and their collaboration with cDCs.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

Adaptive Th1 responses are important for resolving intracellular bacterial infections such as those caused by Salmonella and Mycobacteria. Priming of CD4+ T cells occurs within the T-cell zones (T zones) of secondary lymphoid tissues and requires cognate interaction between dendritic cells (DCs) and naive CD4+ T cells 1. After priming, T cells upregulate CD69 and CD44 and downregulate L-selectin (CD62L) and begin to proliferate. These events occur rapidly after Salmonella Typhimurium infection (STm) 2 and are detectable within the first 24 h. In parallel, T cells can acquire Th1 features such as the capacity to produce IFN-γ 3. In the absence of Th1 differentiation and IFN-γ production, clearance of STm infections is markedly impaired and infection is more disseminated 4–10.

DCs are the most potent APCs. As immature cells, DCs are strategically located in non-lymphoid tissues where they are likely to encounter antigen. After antigen encounter, DCs migrate to the T zones of secondary lymphoid tissues to present it to naive T cells. In secondary lymphoid tissues, in the steady state, several populations of resident DCs can be found and the role of these cells in priming T-cell responses has been studied 11, 12. Importantly, during infection or inflammation, another population of DCs differentiate from recruited blood monocytes. 13–16. These cells, monocyte-derived DCs (moDCs), are characterized by lower expression of CD11c than resident, conventional DCs (cDCs), yet they maintain monocyte markers such as CD115, Ly-6C and CD11b. This population has been described in several infection models, and despite this common phenotype, the role of moDCs in each infection is tailored to the specific threat. For instance, after Listeria monocytogenes infection, a TNF/iNOS-producing DC subset (TipDCs), is important for the control of infection in a TNF-α-dependent manner, but do not contribute to T-cell priming 17. In contrast, during responses to Leishmania18 and influenza 19, 20, DCs expressing monocyte markers are called inflammatory DCs, are important sources of IL-12 and are directly involved in Th1 priming. Despite reports conferring different names to such populations, what is clear is that in each case the surface phenotype of these populations is consistent within infections and they have a monocytic origin 13, 14. Therefore, multiple DC populations can be present in the T zone and participate in T-cell priming 21–23. During STm infection, a number of additional cellular subsets have been observed. One of these, expressing CD11cintCD11b+TNF-α+iNOS+, is found to be present by the third day of infection in mucosal and systemic lymphoid tissues. Nevertheless, despite the expression of DC markers, these cells were not found to contribute to T-cell priming but did augment bacterial killing 24, 25. Thus, how Th1 responses to STm develop is unresolved.

In this study, we show that moDCs accumulate in the T zone of responding lymphoid tissues within 24 h of STm infection and this was dependent upon bacterial viability rather than virulence. moDCs produce TNF-α and are required to prime but not sustain Th1 responses. Significantly, moDCs were able to induce T-cell proliferation ex vivo without further antigen exposure and this was largely TNF-α-dependent. Furthermore, moDCs synergize with cDCs to augment Th1 priming. Thus, a key mechanism that drives efficient Th1 priming and IFN-γ production in response to STm infection is the involvement of moDCs and their co-operation with cDCs.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

F4/80+CD11c+ DCs are present in the T zone by 24 h after STm infection

In earlier studies 6, 26, we observed F4/80+ cells in the T zones of STm infected but not naive mice. In the current study, we assessed their appearance and function in detail. Immunohistology showed that F4/80+ cells accumulate in the T zones of spleens 24 h after STm infection but not in naive mice nor after immunization with the STm flagellin protein (FliC) or alum-precipitated proteins (Fig. 1A). To further characterize these T zone-localized cells, we used confocal microscopy. While in the red pulp of the spleen, F4/80+ cells are overwhelmingly CD11c in the T zone, >99% of T zone F4/80+ cells were also CD11c+ (Fig. 1B). This was further supported by positive staining of DCs for GR1 and Ly6C (Fig. 1B and Supporting Information 1).

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Figure 1. STm infection induces a rapid accumulation of F4/80+CD11c+ cells into the T zone. (A) Immunohistological assessment of F4/80 (blue) and IgD (brown) expression on spleen sections from non-immunized mice or mice immunized i.p. with 5×105 STm, 20 μg FliC or NP-CGG/Alum for 24 h. Left column scale bar indicates 200 μm. The right column is a magnification of the boxed T zone areas (40×); scale bar indicates 25 μm. Arrows indicate F4/80+ cells in the T zone. (B) Composite confocal representative images of spleen cryosections 24 h after STm infection, showing F4/80 or GR1 (red), CD11c (blue) and IgM (gray). The left hand column shows low-magnification images of white and red pulp to identify the T zone; scale bar represents 200 μm. The centre and right columns show higher magnification images (63×) of the white boxed T zones for the expression of F4/80 CD11c and GR1 CD11c; scale bar represents 25 μm. (C) Representative polychromatic dotplots of splenocyte cell suspensions from non-immunized mice or mice infected with 5×105 STm for 24 h. Expression of CD11c (blue) and F4/80 (red) was analyzed. Dot plots show the F4/80+ gate, which is selected to further illustrate the expression of CD11c and CD11b within those cells in the dotplots to the right. Gate and quadrant frequencies are stated. (D) Dot plots evaluating the expression of CD11c and CD11b show cDCs (CD11c+CD11b+, blue) and moDC (CD11c+CD11bhi, pink) in the STm-infected mice (a population of cells CD11b+CD11c are also shown and they represent monocytes and granulocytes). Gate frequencies are stated in red. Their phenotype is further analyzed by the expression of F4/80 and GR1, CD115 and MHC-II; in this case, cells with the cDC gate (blue) and moDC gate (pink) are overlaid in each dot plot. Histograms to the right show phenotype characterization of each population (shaded histogram shows isotype control). (E) Frequency of bacterial infection in cDCs or moDCs sorted (gated as in Fig. 1D) from WT mice infected for 24 h (top graph) and numbers of splenic GFP+ cDCs or moDCs from non-infected or infected mice with STm for 24 h (lower graph and flow cytometry panels, frequencies are stated). *p≤0.05 by the Mann–Whitney test. Experiments were repeated three times with at least four mice per group.

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To characterize this population further, we used multicolour flow cytometry. A polychromatic dot plot shows an increase of CD11c+F4/80+ cells after infection (pink and purple cells), supporting the confocal studies. Further analysis of F4/80+ cells showed that the majority also express high levels of CD11b (Fig. 1C). Detailed characterization of the CD11c+ populations from naive and 24 h STm-infected spleens showed that after infection, there was an increase of CD11cintCD11bhi cells that were GR1+, F4/80+, CD115+ and MHCII+ (Fig. 1D, and Supporting Information Fig. 1B; pink shading/line on dot plot and histogram). This phenotype is consistent with the described phenotype of moDCs and inflammatory DCs 13, 14, 27. The identity of these cells as moDCs and inflammatory DCs was also confirmed by assessing the expression of CD11b, Ly-6C and MHC-II (MHC class II) (Supportive Information 1A). This showed that when the CD11bhiLy6C+MHC-II+ population, only observed after STm infection, was backgated to assess their CD11c and CD11b expression, they corresponded to the population we observed and characterized as CD11cintCD11bhiF4/80+GR1+. For consistency, we refer to this population as moDCs throughout. Neither cDCs nor moDCs cells expressed CD3, CD19, DX5 (used as exclusion markers) or CD138 (data not shown). Similar results were found in mouse strains other than C57BL/6 such as Balb/c.

We also addressed the level of infection in cDCs and moDCs by examining bacterial carriage in these populations by two methods. To do this, we infected mice with STm for 24 h before cell sorting the cells into cDC and moDC populations and assessing bacterial numbers by direct culture (Fig. 1E). In addition, we also infected mice for 24 h with STm that constitutively express GFP (STmGFP) and looked for GFP expression within cDCs and moDCs. As shown in Fig. 1E, a higher proportion and number of moDCss were GFP+ compared with cDCs.

Accumulation of moDCs is most dependent upon bacterial viability

We next assessed the features associated with the accumulation of moDCs by giving different bacterial strains or bacterial antigens and examining moDC numbers in the spleen 24 h later. The induction of moDCs was independent of virulence since infection with similar numbers of attenuated or virulent STm (attenuated through two independent mechanisms, see Materials and methods) induced similar levels of moDC accumulation (Fig. 2A). Furthermore, the induction was most dependent upon bacterial viability since immunization with heat-killed (hk) bacteria or soluble FliC or LPS resulted in substantially fewer moDC being detectable (Fig. 2A). In contrast, after all antigens cDC numbers were similar 24 h after administration (Fig. 2B). Thus, viability of the bacterium, rather than its virulence or its components, is most important for inducing the greatest increase in moDC number. The accumulation of moDCs after STm was not solely restricted to the spleen since mice infected i.p. or s.c. for 24 h had increased moDC numbers in the lymphoid organ draining the site of infection (Fig. 2C).

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Figure 2. MoDCs produce TNFα and their accumulation requires bacterial viability. Quantification of (A) moDCs and (B) cDCs in murine spleens 24 h after i.p. immunization with STm strains SL3261, SL1344, SL1344 SPI2 mutant, FliC, LPS or hk SL3261. N.I., non-immunized. (C) Evaluation of the number of moDCs 24 h after i.p. or s.c. immunization. (D) Costimulatory marker expression in non-immunized mice and 6, 18 and 24 h after infection. Histograms show the expression of CD86 and CD40 on cDCs and moDCs during the time course. The far right histogram shows the CD86 expression 24 h after infection with STmGFP on cDCs, moDCs and moDCs in which GFP was detected. (E) Intracellular IL-12 and TNF-α expression 18 h after STm infection. Spleens were harvested and cell suspensions were depleted using CD19, CD5 and DX5 MACS beads. Cells were cultured for 4 h at 37°C. Intracellular cytokines were assessed by flow cytometry, a relevant PE-Isotype control was used in all samples with no significant staining detected. Graphs show IL-12 and TNFα production from splenic cDCs and moDCs from non-infected mice or mice infected for up to 48 h. Experiments were repeated three times with at least four mice per group. Statistics show p values relative to non-immunized controls; NS, not significant; *p≤0.05, **p≤0.01.

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Analysis of costimulatory molecule expression revealed that moDCs upregulate CD86 and CD40 by 6 h after infection (Fig. 2D), though the kinetics of this was marginally slower than that of cDCs. Infection with STmGFP for 24 h showed that GFP+ moDCs had the highest expression of CD86. We then examined cytokine (TNF-α and IL-12) production by cDCs and moDCs at 18 h (peak expression levels) by intracellular staining. As expected, after STm infection cDCs produced IL-12 28, while moDCs were the main source of early TNF-α and this cytokine profile was maintained throughout the first 48 h of infection (Fig. 2E). Expression of iNOS by moDCs was not detected by intracellular staining (data not shown). The results show that moDCs and cDCs upregulate costimulatory molecules in the spleen within 24 h of infection and contribute different cytokines to the response.

The absence of moDCs impairs early T-cell activation

To assess the contribution of moDCs to T-cell priming and differentiation, we used clodronate liposomes to deplete macrophages and monocytes 29. Mice were injected i.p. with either clodronate-liposomes or PBS-liposomes 24 h before STm infection. Spleens were then analyzed by confocal microscopy and flow cytometry 24 h after infection when moDCs are present in the T zone (Fig. 1A).

As shown in Fig. 3A by confocal microscopy, treatment with clodronate-liposomes but not PBS-liposomes depleted red pulp macrophages and moDCs. In mice treated with clodronate liposomes, moDC numbers were tenfold lower after infection compared with those in mice treated with PBS liposomes (Fig. 3B). In contrast, although there was some reduction (30% median fall) in cDC numbers after clodronate depletion, this difference did not reach significance. Furthermore, confocal microscopy confirmed the presence of cDCs in the T zones of both groups of infected mice (Fig. 3B). Depletion of moDCs resulted in an impaired capacity to prime CD4+ T cells after STm as nearly tenfold fewer CD69+ T cells were detected (Fig. 3C, left graph). In contrast, in mice immunized with hk STm, which results in lower levels of moDCs (Fig. 2A), there was no difference in CD69 expression on T cells (Fig. 3C right graph). Therefore, the use of clodronate-liposomes before infection prevents the accumulation of moDCs in the T zone and this results in impaired CD4+ T-cell priming.

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Figure 3. Treatment with clodronate liposomes prevents accumulation of moDCs into splenic T zones after STm infection. Mice were treated i.p. with PBS or clodronate liposomes (200 μL) 24 h before STm infection. (A) Representative composite, confocal images of spleen sections 24 h after STm infection stained for F4/80 (red), CD11c (blue) and IgM (gray). The left column shows spleen sections from PBS-liposome treated mice and the right column shows sections from clodronate-liposome treated mice. Low-magnification (10×) images identify T zones (scale bar=200 μm) and arrows show higher magnification (63×) of the white boxed areas (scale bar=25 μm). (B) Graphs show the absolute numbers of splenic moDCs (left) and cDCs (right) in non-immunized (N.I.) and in STm-infected mice treated with either PBS or clodronate liposomes. (C) Quantification of CD69+ T cells in the spleen before and after STm (left) or hk STm (right) administration in mice treated with PBS or clodronate liposomes. Experiments were repeated four times with four mice per group. NS, not significant; *p≤0.05; **p≤0.01 (Mann–Whitney test).

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Impaired T-cell activation in the absence of moDCs diminishes IFN-γ production from CD4+ T cells

We next studied what effects depleting moDCs had on T-cell differentiation. Mice were treated with either clodronate or PBS liposomes 24 h before STm-infection and then during infection to maintain depletion. A week after infection, intracellular IFN-γ expression in CD4 T cells was evaluated by ex vivo restimulation. As shown in Fig. 4A, in mice treated with clodronate before STm infection had lower frequencies and numbers of IFN-γ+ T cells compared with PBS-treated STm-infected mice. This lower IFN-γ response was not due to differences in bacterial numbers since bacterial burdens were similar between the two groups that received liposomes, reflecting the findings found in a previous report 30.

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Figure 4. Absence of moDCs at the time of infection diminishes IFN-γ production by CD4+ T cells to STm but does not affect plasma cell expansion. Mice were treated i.p. with PBS or clodronate liposomes (200 μL) (A) 24 h before (N.I. – non-immunized) or (B) 3 days after STm infection. Infection was allowed to continue until day 7 and during this period liposome treatment was administered every 3 days to maintain cell depletion. At day 7, spleens were harvested and intracellular IFN-γ was assessed in CD4+ T cells following 6 h of in vitro re-stimulation with anti-CD3 (10 μg/mL, precoated well) or with culture medium alone. Intracellular cytokines were analyzed by multicolor flow cytometric analysis and total T cells were gated using CD3. The numbers in the quadrants show the frequency of CD3+CD4+IFN-γ+, with the value of the corresponding cells cultured in medium alone subtracted. Graphs show the absolute number of IFN-γ+ CD4+ T cells or splenic bacterial burdens from mice with and without clodronate treatment. Dotplots are representative of three independent experiments with at least four mice per group. (C) Immunohistological assessment of IgG2a (blue) and IgD (brown) expression on spleen sections from mice treated with clodronate or PBS liposomes and STm infected as in section A; scale bar indicates 200 μm (10×). Flow cytometric plots and histograms show intracellular IgG2a levels. Plasma cells were identified by the expression of CD138 and B220; frequencies are stated in the plots. Plots are representative of three independent experiments with four mice per group. Graphs show the quantification of IgG2a+ plasma cells based on counts from tissue sections (top graph) or the absolute number of IgG2a+ cells per spleen calculated from the flow cytometric data (bottom graph). N.I., non-immunized; NS, not significant; **p≤0.01 (Mann–Whitney).

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We next assessed whether moDCs were required to sustain Th1 cells after T-cell priming by depleting moDCs when T-cell responses were established. WT mice were infected with STm and treated with either clodronate or PBS liposomes 3 days later and treated again 72 h to maintain depletion. After 7 days of infection, intracellular IFN-γ production was assessed by ex vivo restimulation (i.e. 4 days after first depletion). These experiments show that when depletion occurred after infection the intracellular IFN-γ response was similar in both groups of mice. Administering clodronate post-infection had no impact on splenic bacterial burdens (Fig. 4B).

Finally, we assessed whether other Th1-associated features of the anti-STm response were affected by loss of moDCs prior to infection by looking at the numbers of extrafollicular IgG2a switched plasma cells on day 7 after infection. In this infection, the induction of the extrafollicular response is T-independent but isotype switching is T-dependent 31. To do this, mice were treated with clodronate prior to infection and infected (as in Fig. 4A). On day 7, the induction of T-dependent plasmablast switching was assessed by immunohistology and flow cytometry (Fig. 4C). This shows that IgG2a switching was not dependent upon moDCs. Thus, moDCs are required for selective elements of Th1 priming during the initial encounter with CD4+ T cells but are dispensable by day 3 after infection, when T-cell priming is established.

MoDCs are sufficient to drive Th1 priming and can synergize with cDCs

To show that moDCs could function as APCs, we analyzed the capacity of cDCs and moDCs to present antigen to transgenic CD4+ T cells and their capacity to promote IFN-γ production. First, cDCs and moDCs were sorted from spleens 24 h after infection and their moDC phenotype confirmed (Fig. 5A, GR1 shown as an example but cells were also assessed for F4/80 expression). When sorted cDCs or moDCs were cultured with CD5-enriched naive CFSE-labeled SM1 CD4+ T cells (at a 30:1 ratio, T: APC) in the presence of added soluble FliC for 4 days, both cDCs and moDCs could induce T-cell proliferation, although cDCs were more efficient (Fig. 5B). Thus, both cDCs and moDCs can process and present antigen. Next, we assessed whether both populations had acquired antigen in vivo and could present this ex vivo in the absence of further antigen encounter. After infection for 24 h, cDCs and moDCs were sorted as before. In all cases, APCs were cocultured in an 1:30 ratio (T:APC) with CFSE-labeled SM1 CFSE-labeled CD4+ T cells for 4 days. In addition, as both populations are co-localized to the T zone in vivo, we assessed whether their co-culture affected priming by co-culturing equal numbers of cDCs and moDCs (total DCs numbers were the same in all three groups). This showed that both DC populations could induce proliferation in the absence of exogenous antigen but having both DC subsets present augmented proliferation (Fig. 5C). These results suggest that DC subsets can collaborate to drive T-cell proliferation.

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Figure 5. MoDCs prime Th1 development ex vivo in a TNF-α dependent manner. (A) cDCs or moDCs were sorted from spleens of mice infected with STm for 24 h. Cells were sorted based on their expression of CD11c, CD11b, GR1 and F4/80 (cDCs, CD11c+CD11b+F4/80GR1, moDCs, CD11cintCD11bhiF4/80+GR1+). Sorted DCs were co-cultured with CFSE labeled T cells in a 1:30 proportion. (B) SM1 T cells were used as responders and 5 μg/mL of FliC added to the culture to evaluate the antigen presentation capacity of the sorted APCs. (C) Sorted cDCs, moDCs or both were cultured with STm-primed T cells from a WT mice infected 7 days previously in the absence of any further exposure to antigen in a 1:30 proportion. All cultures were maintained for 4 or 5 days. T-cell division was evaluated by CFSE dilution. Histograms show CFSE dilution on CD3+CD4+ gated cells. (D) ELISPOT assay for IFN-γ and IL-4 was performed on cocultures of sorted cDCs (open bars), moDCs (filled bars) or both (dashed bars) with naïve SM1 T cells (1:30 ratio). ELISPOT plate was cultured for 4 days. Graph shows numbers of SPU. Culture of cDCs or moDCs or T cells alone induced no SPU. (E) ELISPOT assay was used to evaluate the IFN-γ production in co-cultures of cDCs or moDCs or both with naïve T cells in medium alone (filled circles) or in the presence of 10 μg/mL of blocking anti-TNFα Ab (open squares). ELISPOT plate was cultured for 4 days at 37°C. NS, not significant; **p≤0.01 (Mann–Whitney). Experiments were independently repeated three times.

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To examine how DC subsets could influence Th1 differentiation, cDCs and moDCs were sorted from spleens of mice infected for 24 h as before. These cells were then cultured with FliC and naive SM1 CD4+ T cells on an ELISPOT precoated plate to evaluate the IFN-γ or IL-4-secretion (Fig. 5D). This showed that moDCs induced greater numbers of IFN-γ producing T cells and fewer IL-4-producing cells than cDCs. Co-culture of T cells with both DC subsets selectively induced greater IFN-γ responses than either component DCs subset, but this was not seen for IL-4 (Fig. 5D). This suggests moDCs are more efficient than cDCs at driving CD4+ T cells to produce IFN-γ but can collaborate with cDCs to augment this. Lastly, in this and other studies 24, moDCs have been identified as major producers of TNF-α. To assess whether this cytokine influenced the priming of IFN-γ-producing cells, we cultured cDCs or moDCs with SM1 T cells in the presence or absence of a TNF-α-neutralizing antibody (Fig. 5E). These experiments show that neutralizing TNF-α reduces the numbers of IFN-γ-producing cells induced by moDCs but not by cDCs. Surprisingly, neutralizing TNF-α only moderated Th1 development when moDCs were cultured alone with SM1 T cells. This diminution was not seen when moDCs were co-cultured with cDCs (Fig. 5E). Therefore, moDCs can present antigen to CD4+ T cells and promote their differentiation to become IFN-γ-producing T cells.

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

Th1 responses are characterized by the induction of IFN-γ and are essential for clearing intracellular infections such as those caused by STm. Our studies indicate that moDCs accumulate in the T zone after STm infection, have encountered live bacteria, can present antigen to T cells and in their absence Th1 responses are impaired. Finally, our data suggest that moDCs can act in conjunction with cDCs to perform this function.

It is significant that the accumulation of moDCs is dependent upon bacterial viability rather than virulence. This offers some explanation as to why hk STm vaccines induce Th2 features but poor Th1 responses 32. The importance of viability has also been demonstrated for the recruitment of TipDCs in response to L. monocytogenes17. This suggests that inducing moDCs is likely to be a key requisite of Th1-promoting adjuvants and that characterizing moDC induction is likely to provide a measure of their success. Interestingly, other subunit components of the bacterium that act through TLRs, such as FliC, do not induce moDC accumulation to the same degree and this parallels the lack of Th1 response seen to flagellin in vivo 6, 33, 34. We have also observed differential Th1 or Th2 T-cell priming to OVA when presented within the bacterium or as an alum-precipitated protein respectively 35. This highlights that T-cell fate is not necessarily an intrinsic property of the T cell but dependent upon the signals received from DCs during priming.

Bacterial virulence is not an important requirement for driving moDC accumulation since virulent bacteria and bacteria attenuated through two distinct mechanisms, aroA-deletion resulting in histidine auxotrophy and ssaV-deletion resulting in impaired secretion of Salmonella Pathogenicity Island II effectors, all induced moDCs to similar levels 24 h after infection. It was not possible to study the response to the virulent STm strain reliably beyond this time due to the intrinsic susceptibility of C57BL/6 or BALB/c mice to virulent strains of STm. Nevertheless, the similar numbers of moDCs found 24 h after infection with attenuated and virulent strains offers an explanation as to why the virulence of a STm strain for a particular mouse strain does not impede Th1 differentiation 36.

Similar to previous reports 24, 25, 37, we show that moDCs responding to STm are the major synthesizers of TNF-α. This reflects the cytokine profile from TipDCs after L. monocytogenes infection 17. In contrast, cDCs were the primary producers of IL-12, a cytokine required for the persistence of Th1 responses after STm infection but not for their induction 38. There may be some pathogen specificity in the cytokine signature of moDCs since the predominant source of IL-12 after influenza infection was moDCs rather than in cDCs 20.

In studies identifying inflammatory cell recruitment after STm 24, 37, Wick and co-workers found limited potential for CD11cintCD11b+ cells to prime OVA-specific OTII CD4+ T cells after in vitro infection with STm expressing OVA, although these cells could present OVA peptide. The differences in the findings between these studies are likely to primarily reflect methodological differences. In the current study, moDCs were crucial in the first 24 h after infection. Their early presence is important since depleting monocytes in vivo before infection impaired Th1 priming, whereas depleting from 72 h after infection did not. These differences were not due to depletion-induced changes in bacterial burdens. In a previous report on the use of clodronate before and during STm infection, the effects of clodronate depletion depended upon the virulence of the strain used. Thus, in an attenuated strain similar to that used in most experiments here, bacterial colonization was not affected by depletion, whereas infection with a virulent strain of STm was affected by clodronate treatment so that mice actually had improved survival and lower bacterial colonization after clodronate treatment 30. Direct involvement of moDCs in priming is shown using sorted moDCs. Using naive, transgenic FliC-specific CD4+ T cells, we show that moDCs can drive IFN-γ production and this was abrogated after neutralization of TNF-α. The effects of TNF-α neutralization on diminishing moDCs-mediated Th1 priming were only apparent when moDCs were cultured on their own with T cells and not when they were co-cultured with cDCs. This effect is striking, although the reason for it is unclear. It suggests that there is some compensatory mechanism for the loss of TNF-α at play when different DC subsets are cultured together with T cells. The details of this cellular collaboration and its mechanism are currently under investigation.

In vivo depletion of monocytes by clodronate prevented the accumulation of moDCs in the T zones of infected mice and markedly reduced early T-cell activation and IFN-γ production. While cells with a similar phenotype to the moDCs described here have been found after immunization with alum-precipitated proteins 39, 40, these cells were found to be located in the medulla of the lymph node and not in the T zone 40. Critically, moDCs were required at the earliest stages of infection, since depletion from the third day did not affect IFN-γ production. These multiple lines of evidence indicate that moDCs are the important drivers of early Th1 responses after STm infection.

Using clodronate liposomes as a method to deplete moDCs has some disadvantages, including one of specificity, since macrophages are also depleted. To further this work in the future, other systems such as using Ccr2−/− mice would help identify how the absence of moDCs impacts Th1 polarization and bacterial clearance 20, 41. The role of moDCs in other infections has been addressed using such a strategy and the results from those studies support our findings on the importance of these cells at the time of priming. However, elegant experiments using CCR2-DTR mice show that in selective fungal infections the depletion of moDCs 2 days after infection can affect T-cell polarization 42. These results might reflect differences between infections, for instance in terms of the kinetics of antigen processing and presentation, but could also suggest that the level and timing of crosstalk between moDCs and cDCs could be different as they observed no difference in T-cell expansion. Lastly, there may be some influence of the pathogen on the host. These possibilities are not mutually exclusive.

Optimal Th1 responses in moDCs cultured with T cells required the presence of cDCs. Such collaboration has been described before in responses to other pathogens 43 and is probably required to ensure the appropriate direction of T-cell polarization. How this collaboration works shows some specificity to the pathogen. Thus, in responses to attenuated yeast the moDCs transfer antigen to cDCs and it is the cDCs that prime T-cell responses 43, whereas in the response to Aspergillus moDCs can present antigen 41. This, in conjunction with the finding that the cytokine profile of these cells is also pathogen-specific 17, 18, 20, 24, highlights the complexity of initiating the adaptive response, and emphasizes a major conclusion from this and similar studies, that the immune response is tailored to the individual pathogen. It is apparent from the current study, using STm, that further analysis need to be done in order to establish how the cDC and moDC populations interact to enhance T-cell responses.

In conclusion, this work describes the early requirement of moDCs for optimal CD4+ T-cell priming and IFN-γ production in response to STm infection. The finding that Th1 responses require the collaboration between multiple DC subsets of discrete origins highlights the importance of studying DC subsets to understand how beneficial and harmful T-cell responses develop in vivo.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

Mice and antibodies

C57BL/6 mice, 6–8 wk, were from Harlan Sprague-Dawley. SM1 2 and TCRβ/δ−/− mice were maintained in-house. Animal procedures were performed with local ethical approval and the UK Home Office (Project license 40/2904) under the Animals (Scientific procedures) Act 1986. Antibodies are listed in Supporting Information Table 1.

Bacterial strains and immunizations

STm SL3261 is an AroA attenuated strain 44. SL1344 is a virulent strain and the SL1344 SPI2 mutant, TL64, lacks ssaV 45. STmGFP was generated as described previously 35, by inserting the eGFP gene via ndeI and xhoI restriction sites into the pettac plasmid, which has a modified tac promoter to enable constitutive gene expression. Mice were infected i.p. with 5×105 live STm. Bacteria were heat-killed by heating at 70°C for 1 h with killing confirmed by culture. Some mice received 20 μg recombinant FliC 6 or 15 μg TLR-grade LPS (Alexis Biochemicals). Tissue bacterial burdens were evaluated by direct culturing.

Immunohistochemistry and confocal microscopy

Immunohistology was performed as described previously 6. Cryosections were incubated with primary unlabeled Abs for 45 min at RT before addition of either HRP-conjugated or biotin-conjugated secondary antibodies and ABComplex alkaline phosphatase (Dako). Signal was detected as described 6.

Confocal staining was performed in PBS containing 10% FCS, 0.1% sodium azide. Sections were mounted in 2.5% 1,4-diazabicyclo(2,2,2)octane (pH 8.6) in 90% glycerol/PBS. Primary Abs were incubated for 1 h at RT, and secondary Abs for 30 min at RT. Confocal images were acquired using a Zeiss LSM510 laser scanning confocal microscope. Signals obtained from lasers were scanned separately and stored in four nonoverlapping channels as pixel digital arrays of 2048×2048 (when taken with the 10× objective) or 1024×1024 (when taken with the 63× objective).

Cell preparation and flow cytometry

Spleens were disrupted and digested with collagenase IV 400 U/mL (25 min at 37°C; Worthington Biochemical). EDTA (5 mM final concentration) was added to stop the reaction. Cells were filtered through a 70-μm cell strainer. DCs were enriched by negative selection using MACS beads and LS columns (Miltenyi Biotec; CD19, CD5 and DX5 beads) and kept in MACS buffer (PBS, 0.5% BSA, 2 mM EDTA) during enrichment (purity ≥75%). Cells were then processed for multicolor FACS analysis with prior blocking with anti-CD16/32 antibody. Primary mAbs or isotype controls were added for 20 min at 4°C and cells analyzed (FACSCalibur cytometer and FlowJo software version 8.8.6).

Intracellular cytokine staining

Intracellular cytokines were evaluated on purified DCs. Enriched DCs (3×106 cells/mL) were cultured for 4 h, with Brefeldin A (BFA, 10 μg/mL) for the last 2 h. Surface staining was performed followed by intracellular staining using standard methods (BD Biosciences). For intracellular IFN-γ staining, T cells were plated at 6×106 cells/mL with 1 μg/mL anti-CD28 Ab and restimulated with 10 μg/mL anti-CD3 or medium for 6 h at 37°C, with Brefeldin A (10 μg/mL) for the last 2 h. After incubation, cells were processed for multicolour FACS analysis. For intracellular Ig Ab staining, splenocytes were processed as above.

Clodronate-liposome treatment of mice

Clodronate (Cl2MDP) liposomes or PBS liposomes (200 μL i.p.) 29 a kind gift from Roche Diagnostics GmbH, were injected 1 day before or 3 days after infection.

In vitro co-culture for T-cell priming

TCRβδ−/− mice were infected (5×105 STm) for 24 h and cell suspensions made using Collagenase IV digestion. Cells were pre-enriched by depleting CD19+ and DX5+ cells using MACS beads before staining with CD11c, CD11b and F4/80 to FACS-sort cDCs (CD11chiCD11b+F4/80) and moDCs (CD11c+CD11bhiF4/80+; purity ≥95%). T cells were obtained from SM1 mice, MACS-enriched (CD5+ selection) and CFSE labeled. DCs were added in a 1:30 proportion (APC:T) and incubated for 4 days before analysis by flow cytometry.

ELISPOT analysis of cytokine secretion

ELISPOT assay for IFN-γ and IL-4 was performed as described before 33 using XMG 1.2 as capture Ab for IFN-γ and a mouse IL-4 ELISPOT kit (eBioscience). Plates (Millipore) were pre-coated overnight at 4°C with capture Ab before adding 3×105 MACS-enriched SM1 T cells. Sorted cDCs or moDCs were used as stimulators in a 1:30 (DCs:T cell) proportion. In cDCs and moDCs co-culture experiments equal numbers of cDCs and moDCs were added to T cells to keep a 1:30 proportion. Cells where restimulated with 5 μg/mL FliC or medium alone with anti-CD28 antibody (1 μg/mL) and cultured for 3 or 4 days at 37°C before adding the detection Ab. The reaction developed using DAB. Spots were counted using the AID ELISPOT Reader System. Counts were expressed as SPUs/5×105 splenocytes.

Statistical analysis

Statistics were calculated using the nonparametric Mann–Whitney sum of ranks test using the Analyze-It program. p values of ≤0.05 were accepted as significant.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

This work was supported by a BBSRC New Investigator Award to AFC. The authors are grateful to the Birmingham Biomedical Services Unit for their technical assistance and to Roger Bird for cell sorting. The authors also thank Robert Kingsley and Gordon Dougan at the Sanger Centre, Cambridge for supplying the Salmonella mutant TL64.

Conflict of interest: The authors declare no financial or commercial conflict of interest.

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  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Materials and methods
  7. Acknowledgements
  8. References
  9. Supporting Information

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