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. Author manuscript; available in PMC: 2018 Feb 21.
Published in final edited form as: Immunity. 2017 Feb 9;46(2):205–219. doi: 10.1016/j.immuni.2017.01.003

CD8+ T cells orchestrate pDC – XCR1+ dendritic cell spatial and functional cooperativity to optimize priming

A Brewitz 1,13, S Eickhoff 1,13, S Dähling 1, T Quast 2, S Bedoui 3, R A Kroczek 4, C Kurts 1, N Garbi 1, W Barchet 5, M Iannacone 6, F Klauschen 7, W Kolanus 2, T Kaisho 8,9,10, M Colonna 11, R N Germain 12, W Kastenmüller 1
PMCID: PMC5362251  NIHMSID: NIHMS851708  PMID: 28190711

Summary

Adaptive cellular immunity is initiated by antigen-specific interactions between T lymphocytes and dendritic cells (DC). Plasmacytoid DC (pDC) support antiviral immunity by linking innate and adaptive immune responses. Here we examined pDC spatiotemporal dynamics during viral infection to uncover when, where and how they exert their functions. We found that pDC accumulated at sites of CD8+ T cell antigen-driven activation in a CCR5-dependent fashion. Furthermore, activated CD8+ T cells orchestrated the local recruitment of lymph node resident XCR1 chemokine receptor-expressing DC via secretion of the XCL1 chemokine. Functionally, this CD8+ T cell mediated reorganization of the local DC network allowed for the interaction and cooperation of pDC and XCR1+ DC, thereby optimizing XCR1+ DC maturation and cross-presentation. These data support a model in which CD8+ T cells upon activation create their own optimal priming microenvironment by recruiting additional DC subsets to the site of initial antigen recognition.

Graphical Abstract

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Introduction

The generation of an adaptive immune response involves multiple myeloid and lymphoid cell populations that act in a highly orchestrated manner to permit optimal cellular interaction and communication (Qi et al., 2014). The critical interaction that initiates an adaptive cell-mediated immune response occurs between antigen-bearing dendritic cells (DC) and T cells, leading to proliferation and differentiation of the latter. Key inputs driving such responses involve signaling through the T cell receptor (signal 1), costimulatory receptors (signal 2), and receptors for inflammatory cytokines (signal 3), along with the actions of chemokines and other chemoattractants that fine-tune the localization of lymphocytes and DC within secondary lymphoid organs (Chen and Flies, 2013; Curtsinger and Mescher, 2010; Fooksman et al., 2010; Thelen and Stein, 2008).

Besides the interaction between antigen-bearing dendritic cells (DC) and T cells other cell types contribute to optimal cell-mediated responses (Bendelac et al., 2007; Martin-Fontecha et al., 2004; Veiga-Parga et al., 2013). Chief amongst these additional players are plasmacytoid DC (pDC) (Yoneyama et al., 2005). pDC morphologically resemble lymphoid cells rather than myeloid cells and on a single cell level are known for their ability to produce high amounts of interferon (IFN) type I (Swiecki and Colonna, 2015). For this function, expression of toll-like receptor-7 (TLR7) and TLR9 located in endosomal compartments of pDC are critical. This allows them to sense viral infections irrespective of viral replication within the pDC.

pDC activity enhances adaptive antiviral CD8+ T cell responses and also contributes to innate host defense by inhibiting viral replication during both, acute and chronic viral infections (Cervantes-Barragan et al., 2012; Swiecki et al., 2010). Currently, it is unclear how pDC exert these dichotomous functions (Reizis et al., 2011a). While they are the most prodigious producers of IFN I on a per cell basis, various other cell types, especially macrophages can produce IFN I upon viral infection. Indeed, the total amount of IFN I in the serum of mice in various experimental systems is unaltered in the presence or absence of pDC (Barchet et al., 2005; Xu et al., 2015). Moreover, pDC appear to be only a transient source of IFN I within the first few days of an infection, with the cells then either being eliminated or inhibited via an IFN I feedback loop (Swiecki et al., 2011). Some data indicate that pDC may support anti-microbial resistance by factors other than IFN I (Ang et al., 2010), while other studies suggest that pDC can contribute to adaptive immunity by acquiring, processing and presenting antigens directly to T cells (Di Pucchio et al., 2008; Mouries et al., 2008; Villadangos and Young, 2008). Nevertheless, the majority of published work supports the notion that the primary effector function of pDC lays within their capacity to produce IFN I (Haeryfar, 2005; Reizis et al., 2011b).

Given that many other leukocytes can produce IFN I, the question then arises as to what the special role of pDC IFN I production might be? One possibility could be the ability to produce large amounts of IFN I very rapidly or the capacity to secrete a wider array of IFN-α (Izaguirre et al., 2003). However, using vesicular stomatitis virus (VSV) as model, subcapsular macrophages and not pDC proved to be the critical early IFN I source involved in host protection (Iannacone et al., 2010). Therefore, we hypothesized that besides the timing or amount of IFN I production by pDC, their motility and capacity for relocalization might allow them to deliver IFN I at specific sites to promote host defense in a unique manner (Asselin-Paturel et al., 2005). This notion is in line with the numerous chemokine receptors expressed on pDC that could direct their migrational pattern during infection (Miller et al., 2012).

Given this idea, we felt it important to characterize the spatiotemporal dynamics of pDC during an ongoing immune response. To this end we analyzed pDC migration patterns in the LN using dynamic intravital 2-photon microscopy (IVM). We found that upon viral infection, pDC either migrated to infected macrophages residing in the subcapsular sinus (SCS) area in a CXCR3-dependent manner or to CD8+ T cell priming sites in a strictly CCR5-dependent manner. We show that pDC were recruited to sites of functional MHCI antigen presentation by CCL3 and CCL4 chemokines that were produced in the context of productive conventional DC (cDC) – CD8+ T cell interactions. Additionally, CD8+ T cell activation led to a rapid production of XCL1, which in turn attracted LN-resident XCR1+ DC to CD8+ T cell priming sites. The co-presence of pDC, XCR1+ DC and CD8+ T cells within a confined microenvironment allowed for optimal signal exchange, in particular involving pDC-derived IFN I. This pDC-derived cytokine optimized the maturation of and cross-presentation by XCR1+ DC, enhancing the developing CD8+ T cell response. These data reveal a biological concept in which T cells actively orchestrate the local development of an optimal priming environment upon their initial successful identification of an antigen-bearing cDC.

Results

pDC maximize CD8+ T cell responses

To understand how pDC contribute to antiviral immune responses we used modified vaccinia virus Ankara (MVA) to study CD8+ T cell responses in the presence or absence of pDC. This attenuated cytopathic virus strain does not replicate in vivo and therefore induces a highly synchronized immune response in space and time. Additionally, when manipulating the IFN I system secondary effects on viral replication and thus antigen abundance that impacts the ensuing CD8+ T cell response may be neglected in this experimental approach. Clec4C is highly expressed by pDC and Clec4c+/DTR mice provide a means to selectively deplete pDC by administration of Diphtheria toxin (DTX) to the animals (Swiecki et al., 2010) (Figure S1A–D). After footpad infection, we found a significant reduction in both the relative and absolute numbers of antigen-specific CD8+ T cells directed against the immuno-dominant epitope B8R20 in the absence of pDC (Figure 1A–C). Among the smaller cohort of activated, antigen-specific CD8+ T cells present on d8 post infection in the DTX-treated animals, we only detected modest alterations in CD8+ T cell differentiation using standard markers for various effector subsets (KLRG1 and CD127) (Figure 1D, 1E). Functionally, there were reduced numbers of IFN-γ-producing CD8+ T cells following in vitro restimulation with B8R20 peptide (Figure 1F–H). Among those IFN-γ-producing CD8+ T cells, there were no significant changes in polyfunctionality (cells additionally producing TNFα and/or IL-2) (Figure 1I, 1J). These data indicate that, in accord with earlier studies, pDC help to promote the magnitude of the CD8+ T cell cohort responding to viral infection, while having a limited impact on the differentiation and functionality of the reduced number of activated T cells.

Figure 1. pDC maximize CD8+ T cell responses.

Figure 1

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(A–E) Analysis of B8R20 specific CD8+ T cell responses d8 p.i. (MVA OVA f.p.), comparing WT and Clec4c+/DTR animals treated with DTX (0.1 μg/day; d-2 to d3), showing (A) representative B8R20 multimer staining, (B) relative and (C) absolute numbers of CD8+ T cells specific for B8R20. (D, E) Relative distribution of B8R20-specific memory subsets indicated by the markers KLRG1 and CD127, shown as representative original plots (D) and as relative numbers (E). (F–J) Cytokine production of CD8+ T cells upon in vitro restimulation with B8R20 peptide showing (F) representative plots, (G) relative and (H) absolute numbers of IFN-γ-producing CD8+ T cells. (I, J) Polyfunctionality of CD8+ T cells (gated on IFN-γ+) shown as (I) representative plot or (J) relative numbers. TE – terminal effector cells, DN – double negative cells, MP – memory precursor cells, DP – double positive cells. Data are shown as mean ± standard deviation and are representative of four independent experiments analyzing at least 3 mice per group. *** = p≤0.001, * = p≤0.05. See also Figure S1.

pDC are recruited to infected macrophages and CD8+ T cell priming sites

To examine whether the specific localization of pDC contributes to these effects on CD8+ T cell responses and if so, how such behavior compares to the activity and positioning of pDC involved in mediating innate host defense, we employed SiglechGFP/GFP mice that have fluorescently marked pDC (Swiecki et al., 2010). IVM was used to visualize and track these cells in the steady state and after infection in the LN. In these mice GFP is also expressed in medullary macrophages of the LN, however these cells can be easily discriminated from pDC based on the brightness of the GFP signal (macrophages are dim) and the morphology of the cells (Figure S2A). The MVA used for infection encoded the protein antigen ovalbumin (OVA), allowing us to use OT-I T cell receptor (TCR) transgenic OVA-specific CD8+ T cells to follow the development of an adaptive immune response at the same time (Kastenmuller et al., 2013b).

In the steady state pDC were distributed throughout the LN paracortex and the interfollicular area (Figure 2A) with preference around high endothelial venules, similar to human tissue (Colonna et al., 2004). Morphologically, pDC resembled lymphocytes and migrated actively at an average speed of 5μm/min (Figure 2B, 2C and Movie S1). Track length and displacement was significantly lower compared to naïve CD8+ T cells (Figure 2D, 2E), indicating that pDC show less extensive tissue scanning as compared to T lymphocytes in LN.

Figure 2. pDC are recruited to infected macrophages and CD8+ T cell priming sites.

Figure 2

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(A) Immunofluorescent (IF) images of the popliteal lymph node (pLN) from a SiglechGFP/GFP mouse showing the distribution of pDC in the steady state. (B) Image from intravital microscopy (IVM), see also Movie S1. (C–E) Steady state migration analysis of pDC and OT-I T cells in the interfollicular area showing the speed (C), translated tracks (D) and track displacement (E). (F) IF images of the subcapsular sinus (SCS) area 10hrs p.i. (G) Spatial frequency distribution of pDC in the LN before and after infection. (H) Image from IVM showing pDC and infected macrophages 9hrs p.i., see also Movie S2. (I) Migration analysis of pDC at the SCS showing speed (I), translated tracks (J) and track displacement (K) p.i. (L) IF images of the LN paracortex 10hrs p.i. showing OT-I T cell and pDC co-cluster. (M) Image from IVM showing pDC co-clustering with OT-I T cells, see also Movie S3. (N–P) Migration analysis of pDC and OT-I co-cluster 9hrs after infection showing the speed (N), translated tracks (O) and track displacement (P). (Q) Images from IVM showing pDC migration towards an OT-I cluster, see also Movie S2. Mice were infected with MVA OVA or MVA OVA tdTomato. Data are representative of more than four independent experiments analyzing at least 3 mice per group. Red bars indicate mean values. Scale bars: 200 μm (A), 50μm (B, H), 30μm (F, M), 20μm (L), 15μm (Q). Data are shown as mean ± standard deviation. *** = p≤0.001. See also Figure S2, Movie S1, S2 and S3.

Eight hours after infection we found that pDC had translocated to two distinct areas of the LN. One fraction of pDC accumulated at the SCS close to the virus-infected macrophages (Figure 2F, 2G). Here they arrested and interacted with infected SCS macrophages (Figure 2H–K and Movie S2). The other fraction of pDC was located around OT-I T cell clusters that formed around a virus-infected cDC in the interfollicular area (Figure 2L). These pDC directly interacted with the antigen-engaged OT-I T cells and possibly with the infected cDC, forming a super-cluster of pDC around the initial CD8+ T cell cluster (Figure 2L–P and Movie S3). When imaging pDC behavior early after T cell activation we found evidence of pDC migration towards CD8+ T cell clusters in line with either CCR5-dependent chemotaxis and/or chemokinesis and subsequent pDC retention at T cell cluster sites (Figure 2Q).

The direct association between pDC and OT-I T cells might reflect an antigen-specific interaction. To examine this possibility, we transferred labelled peptide-pulsed (OVA257) splenic cDC and OT-I T cells into pDC reporter mice and visualized their interactions. In this experimental set-up antigen-presentation is restricted to the transferred cDC. Twelve hours after cDC transfer we found tight interactions between peptide-pulsed cDC and OT-I T cells. Importantly, under this condition pDC still formed clusters around OT-I T cells suggesting that pDC – CD8+ T cell communication does not require antigen-specific interactions (Figure S2B).

In summary, we conclude that pDC show a rapid intranodal relocalization during viral infection. Some pDC migrate to the SCS to interact with infected macrophages, while another group of pDC accumulates in the vicinity of CD8+ T cells that are activated by virus-infected, antigen-presenting cDC.

pDC migrate to infected sites via CXCR3 chemokine receptor

Given the relocation of pDC to two distinct sites within infected LN and some evidence of directional migration, we hypothesized that chemokines were playing important roles in promoting these positional changes. Several groups have shown that CXCR3 plays a non-redundant role in controlling lymphocyte migration to the SCS for interactions with infected macrophages (Groom et al., 2012; Kastenmuller et al., 2013b; Sung et al., 2012). Therefore, we speculated that CXCR3 might also be critical for pDC to migrate to this region of the LN. To examine this possibility, we crossed pDC reporter mice to CXCR3-deficient animals and infected these mice with MVA OVA tdTomato. In contrast to WT animals, CXCR3-deficient mice did not show efficient translocation of pDC to the SCS (Figure 3A, 3B and Movie S4). Semi-automated quantification supported these visual interpretations (Figure 3C). The few pDC that were present at the SCS in Cxcr3−/− animals interacted with the infected macrophages leading to migrational arrest similar to their WT counterparts (Figure 3D, 3E). In line with a critical role for CXCR3 for pDC migration to the SCS, we found an upregulation of both CXCL9 and CXCL10 after infection with MVA (Figure 3F, 3G). In the context of published work showing that SCS macrophages are primary targets of viral infection and promote viral replication in the LN, these data suggest that CXCR3-mediated pDC recruitment to infected SCS macrophages could contribute to control local viral replication and gene expression (Iannacone et al., 2010; Swiecki et al., 2010). To directly test this notion, we infected WT, Cle4c+/DTR and CXCR3-deficient animals with replication competent VSV in the footpad and analyzed the viral titers 24hrs later. If pDC were depleted, we did not detect differences in the viral titers in the popliteal LN but there was a significant dissemination of virus to the upstream inguinal LN (Figure S3A). In contrast, in CXCR3-deficient animals this enhanced lymphatic viral dissemination was not observed. However, CXCR3-deficient animals had significantly increased pDC numbers in the LNs before infection as compared to control animals, while the migrational speed was unaltered (Figure S3B, S3C). The increased density of pDC in the CXCR3-deficient mice may place enough of these cells near to the SCS macrophages in the steady state that chemokine-mediated guidance is no longer a limiting feature of the response upon infection. These findings concerning the innate antiviral properties of pDC left open the question of how pDC are differentially recruited to CD8+ T cell priming sites and what possible functions they exert in support of T cell activation and adaptive immune responses.

Figure 3. pDC accumulate at the SCS in an CXCR3-dependent manner.

Figure 3

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(A, B) IF images showing pDC localization 7–10hrs p.i. in (A) WT and (B) Cxcr3−/− mice; dashed line indicates SCS area. (C) Spatial frequency distribution of pDC comparing WT (SiglechGFP/GFP) and Cxcr3−/− (SiglechGFP/GFP) mice 10hrs p.i. (D) Image from IVM showing Cxcr3−/− (SiglechGFP/GFP) pDC 7hrs p.i. (E) Analysis of the mean velocity of pDC in WT (SiglechGFP/GFP) and Cxcr3−/− (SiglechGFP/GFP) mice. (F, G) Quantification of CXCL9 and CXCL10 from WT and Clec4c+/DTR animals treated with DTX using pLN homogenates from mock treated or infected conditions showing dot plots (F) and ELISA (G). Ratios indicate relative intensities adjusted to controls (HSP60). Mice were infected with MVA OVA tdTomato. Data are representative of (A–E) five independent experiments (n≥3), (F) one experiment from pooled samples (n=9) or (G) two independent experiments (n=3). Red bars indicate mean values. Scale bars: 150μm (A, B), 30μm (C). *** = p≤0.001. Please see also Figure S3 and Movie S4.

CCR5 guides pDC to CD8+ T cell priming sites

On the assumption that chemokines also direct pDC towards sites of CD8+ T cell priming by antigen-bearing cDC, we first sought to determine if virus-induced inflammatory signals played a role in this colocalization phenomenon or if the relevant signals arose from the interacting T cells and cDC. For this purpose, we designed an in vivo system that activates CD8+ T cells, yet lacks virus-associated inflammatory cues. Labeled OT-I T cells were transferred into pDC reporter mice that were then immunized with LPS-free OVA. T cell and pDC behavior was then visualized in these animals. Similar to what we observed during viral infection, pDC colocalized with the arrested OT-I T cells, forming large cellular aggregates (Figure S4A, S4B). This indicated that T cell- or cDC-derived rather than virus-promoted signals might lead to pDC recruitment to T cell priming sites. Besides CXCR3, CCR5 is highly and homogenously expressed on pDC and plays an important role for pDC transmigration through inflamed vessels (Diacovo et al., 2005; Miller et al., 2012). CCR5 interacts with CCL3, CCL4 and CCL5, chemokines that are produced by activated T cells (Dorner et al., 2002) and that prior studies showed played a key role in cell – cell interactions involved in the generation of cell-mediated immune responses (Castellino et al., 2006; Hugues et al., 2007). Indeed, we found a strong increase in CCL3, CCL4 and CCL5 in LN homogenates 8hrs after infection with MVA OVA (Figure 4A, 4B). To assess the role of CCR5 in intranodal migration of pDC, we crossed CCR5-deficient animals to pDC reporter mice and visualized pDC behavior in vivo upon viral infection using IVM. In CCR5-deficient mice, pDC were not found at OT-I T cell clusters (Figure 4C, 4D and Movie S5). Similarly to MVA infection, we detected a CCR5-dependent accumulation of pDC around OT-I T cell clusters in the context of a low dose VSV OVA infection (Figure S4C–E).

Figure 4. pDC are recruited to CD8+ T cell priming sites via CCR5.

Figure 4

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Dot blot detecting CCL3, CCL4 and CCL5 (A) and ELISA detecting CCL3 (B) using LN homogenates from mock treated and infected (8hrs) animals. Ratio indicates relative intensities normalized to controls (HSP60). (C) Quantification of pDC in proximity to OT-I T cell clusters in WT, Ccr5−/− and Cxcr3−/− animals (all mice SiglechGFP/GFP) 10hrs p.i. (D) IF image from Ccr5−/− (SiglechGFP/GFP) mice showing pDC and OT-I T cells 10hrs p.i. (E) IF image from Cxcr3−/− (SiglechGFP/GFP) mice showing pDC and OT-I T cells 10hrs p.i. (F, G) Image from IVM of pLN showing OT-I T cell cluster and pDC from Ccr5−/− (SiglechGFP/GFP) (F) and Cxcr3−/− (SiglechGFP/GFP) mice (G). (H) CCL3 ELISA from supernatants of activated (αCD3 and αCD28 – 8hrs) or non-activated, purified naïve (CD44low) CD4+ and CD8+ T cells. CD69 indicates extent of T cell activation within cultures. (I–K) IF images and quantification of OT-I T cells and pDC localization 8hrs p.i., comparing isotype- with αCCL3 and αCCL4 treated animals. Mice were infected with MVA OVA. Data are representative of (A) one experiment from pooled samples (n=9), or (C–G and H–K) at least three independent experiments (n=3) or (B, H) two independent experiments (n=3). Scale bars: D (150μm and 15μm), E (150μm and 20μm), F and G (20μm), J (150μm and 10μm), K (200μm and 10μm). Data are shown as mean ± standard deviation. Red bars indicate mean values. *** = p≤0.001, * = p≤0.05, ns = non-significant. Please see also Figure S4, Movie S5 and S6.

When we visualized pDC after MVA infection in CXCR3-deficient mice, focusing on the CD8+ T cell priming sites, we found a significantly enhanced recruitment of pDC that formed groups of up to about 60 cells interacting with the OT-I T cell cluster (Figure 4C, 4E, 4G and Movie S6). However, this enhanced recruitment may be simply based on the increased numbers of pDC present in CXCR3-deficient mice (Figure S3B). The steady state distribution and migrational speed of pDC in skin-draining LNs was unaltered in the absence or presence of CCR5 or CXCR3 (Figure S3C, S4F).

We have previously shown that CD4+ and CD8+ T cells are initially activated separately on distinct cDC (Eickhoff et al., 2015). Therefore, we wanted to address whether pDC are relocalized to and interacted with activated CD4+ T cells and if so whether this process was also CCR5-dependent. To this end we transferred TCR transgenic CD4+ T cells (SMARTA) into pDC reporter mice and infected them with MVA GP (glycoprotein from LCMV). Eight hours after infection we observed CD4+ T cell clusters in the paracortex, in deeper areas of the LN than observed for their CD8+ T cell counterparts. Importantly, we did not detect large pDC superclusters around activated CD4+ T cells, arguing that pDC are more effectively recruited towards activated CD8+ T cells (Figure S4G). However, CD4+ T cell clusters consisted of fewer T cells than CD8+ T cell clusters and some pDC did accumulate adjacent to CD4+ T cell clusters (Figure S4H and S4I). Therefore, quantitative (number of cells producing a chemokine) rather than qualitative differences (selective chemokine production by CD8+ vs. CD4+ T cells) may be the basis for the observed differences. To gain further insight into this issue, we stimulated purified naïve CD4+ and CD8+ T cells in the presence of plate-bound anti-CD3 and anti-CD28 antibodies and probed the supernatant for CCL3 using ELISA. Eight hours after activation we detected a robust amount of CCL3 in the supernatant of CD8+ T cell cultures (Figure 4H). In contrast, we failed to detect CCL3 in CD4+ T cell cultures. Importantly, the extent of activation as measured by CD69 upregulation was comparable in both cultures (Figure 4H). These results are in line with a predominant recruitment of pDC to CD8+ rather than to CD4+ T cell priming sites based on selective production of CCL3 by CD8+ T cells. Prior studies showed that the interaction of antigen-specific CD4+ T cells with antigen-bearing cDC led to local generation of a combination of CCL3 and CCL4, suggesting that T cell induction of chemokine production by cDC could account for the less robust but measurable in vivo recruitment of pDC to antigen-driven CD4+ T cell clusters (Castellino et al., 2006).

To further examine whether CCL3 and CCL4 were the critical chemokines leading to the recruitment of pDC to CD8+ T cell activation sites, we blocked CCL3 and CCL4 signals in vivo using neutralizing antibodies (Castellino et al., 2006) and analyzed the migratory behavior of pDC upon viral infection in the presence of OT-I T cells. Such treatment significantly reduced the numbers of pDC at CD8+ T cell priming sites as compared to what was observed in animals treated with isotype control antibodies (Figure 4I–K). Overall, our results are in line with the interpretation that CCR5 instructs specific recruitment of pDC towards CD8+ T cell priming sites, where CCL3 and CCL4 are secreted by activated CD8+ T cells and likely their interacting cDC.

IFN I signaling involving cDC is critical for optimization of the antiviral CD8+ T cell response

Having established how pDC are recruited to CD8+ T cell priming sites, we next examined how the presence of pDC contributed to enhancing the CD8+ T cell response. The central function of pDC appears to be their capacity to produce high amounts of IFN I upon viral infection (Haeryfar, 2005). However, after depletion of pDC we did not observe a reduction in the total IFNα response in the draining LN 8hrs after infection (Figure 5A). To assess whether IFN I was critical for pDC function in our experimental system, we crossed Clec4c+/DTR mice with Ifnar1−/− animals. When analyzing the immune response on d8 after infection with MVA OVA, we found no significant differences regarding the antigen-specific CD8+ T cell response when comparing Ifnar1−/− animals that did or did not have a pDC population (Figure 5B–E and S5A–D). The loss of the augmenting effect of pDC in Ifnar1−/− mice indicated that IFN I is critical for pDC function in our experimental model. Together with our results that pDC do not substantially alter the total amount of IFNα in the LN (Figure 5A), we conclude that pDC derived IFN I acts locally on the interacting cellular partners. To identify the cell population(s) on which IFN I is acting to optimize CD8+ T cell priming, we first analyzed mice that specifically lack IFNAR1 on T cells (Cd4-cre x Ifnar1flox/flox) and compared them to their WT littermates. In four independent experiments, we consistently observed a slightly reduced antigen-specific CD8+ T cell response among the IFNAR1-negative T cells (Figure 5F, 5H and S5E–H). However, the statistical analyses of the pooled data sets did not show significant differences (Figure 5G, 5I). While these results indicate some contribution of direct IFN I signaling in CD8+ T cells to the effects mediated by pDC, this mechanism can only partially account for the differences we observed when removing the pDC compartment (Figure 1). Therefore, we speculated that pDC-derived IFN I might additionally act on cDC to optimize their function. To address this hypothesis, we analyzed mice that specifically lack IFNAR1 on DC (Itgax-cre x Ifnar1flox/flox). In such animals, there was a substantially reduced CD8+ T cell response on d8 post immunization in comparison to their WT littermates (Figure 5J–M, S5I–L). This result pointed to a critical role of IFN I in optimizing DC functionality.

Figure 5. pDC function and target cells of IFN I.

Figure 5

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(A) IFNα ELISA from LN homogenates from DTX-treated WT and Clec4c+/DTR mice treated 8hrs p.i.. (B–M) Analysis of immune response on d8 p.i., comparing Ifnar−/− and Ifnar−/− x Clec4c+/DTR animals (B–E), CD4 Cre x Ifnarflox/flox with littermates (F–I), CD11c Cre x Ifnarflox/flox with littermates (J–M). (B, F, J) Representative FACS plots of IFNγ-producing CD8+ T cells. (C, G, K) Absolute numbers of IFN-γ-producing CD8+ T cells. (D, H, L) Representative FACS plots showing TNF-α and IL-2 production (gated on CD8+ and IFNγ+). (E, I, M) Quantitative analysis of TNF-α and IL-2 producing CD8+ T cells (gated on CD8+, IFN-γ+). Mice were infected with MVA OVA. Data are representative of (A) three (n=3) independent and (B–M) four (n=4) independent experiments analyzing at least 4 animals per group. (G, I) Show pooled data from four independent experiments (n=16). Data are shown as mean ± standard deviation. ** = p≤0.01, * = p≤0.05. Please see also Figure S5.

pDC locally cooperate with cross-presenting XCR1+ DC via IFN I

Given the physical proximity among pDC, CD8+ T cells and infected cDC within multicellular clusters, we first analyzed the expression of costimulatory molecules on infected cDC in the presence or absence of IFN I signaling. We infected Itgax-cre x Ifnar1flox/flox and control littermates i.v. with MVA GFP and analyzed the expression of CD80, CD86 and CD40 on infected GFP+ cDC 8hrs later. These analyses did not reveal any significant differences in the expression of costimulatory molecules on infected DC in the presence or absence of IFN I signaling (Figure 6A). To test whether the IFN I might act on non-infected cDC, we isolated splenic cDC 36h after infection, identified them by surface marker expression (XCR1 vs. CD11b) and analyzed the expression pattern of costimulatory molecules. This revealed a striking reduction in CD40, CD80 and in particular CD86 expression on XCR1+ DC but not CD11b DC, if IFNAR1 was absent on cDC (Figure 6B, 6C). To determine if this finding was related to pDC function, we depleted pDC, infected the animals and analyzed the expression of costimulatory markers on splenic DC 36h later. Again, we detected a significant reduction of CD80 and CD86 expression on XCR1+ DC in pDC-depleted as compared to non-depleted control animals (Figure 6D, 6E). These data indicated that pDC-derived IFN I acted selectively on XCR1+ DC rather than CD11b+ DC. It is well established that XCR1+ DC are of particular importance for cross-presentation of viral antigens and also serve as a platform that allows interaction with both antigen-specific CD4+ and CD8+ T cells for optimal T cell programming and memory formation in the context of viral infections (Bachem et al., 2012; Eickhoff et al., 2015; Hor et al., 2015). Since MVA does not replicate and therefore heavily depends on cross-presentation for CD8+ T cell priming (Gasteiger et al., 2007), we investigated the role of XCR1+ DC in MVA-induced CD8+ T cell responses. Indeed, the CD8+ T cell response directed against the immunodominant viral epitope (B8R20) was strongly impaired if XCR1+ DC were depleted, confirming the critical role of this DC subset for generating functional CD8+ T cell responses (Figure S6A–H). To further analyze whether XCR1+ DC functionality was enhanced by pDC, we probed the capacity of XCR1+ DC to drive OT-I T cell proliferation in the presence or absence of pDC. As we previously demonstrated, the initial activation of CD8+ T cells after vaccinia virus infection is independent of cross-presenting XCR1+ DC, but is driven by infected DC, predominantly the CD11b DC subset (Eickhoff et al., 2015). To discriminate between these two populations (infected DC vs. cross-presenting XCR1+ DC), we transferred labeled OT-I T cells 60h after infection and analyzed the proliferation profile three days later. At this time point post infection, infected cDC had largely died and been cleared, therefore antigen-presentation is dominated by cross-presenting XCR1+ DC (Eickhoff et al., 2015; Gasteiger et al., 2007). As expected, depletion of XCR1+ DC largely abrogated OT-I proliferation in vivo in this time-frame as compared to control animals (Figure S6I, S6J). In a similar experimental set-up using dye labelled OT-I GFP T cells, pDC depletion also led to a significantly reduced OT-I proliferation in vivo as compared to non-depleted animals (Figure 6F, 6G). To look for a link between these data and the requirement for CCR5 in directing pDC to CD8 priming sites, we analyzed the proliferation of WT OT-I T cells in CCR5-deficient hosts. Similar to pDC-depleted animals, OT-I T cells proliferated significantly less in CCR5-deficient hosts (Figure 6F, 6G). Taken together, these results indicate that pDC-derived IFN I directly or indirectly activates XCR1+ DC to optimize the expression of costimulatory molecules and their ability to activate CD8+ T cells via cross-presentation.

Figure 6. pDC optimize XCR1+ DC maturation and cross-presentation.

Figure 6

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(A) Phenotypic analysis of GFP positive splenic DC and their maturation comparing Ifnar1flox/flox (WT) and CD11c Cre x Ifnar1flox/flox mice 8hrs p.i. (MVA OVA GFP i.v.). (B, C) Representative histograms and quantitative analysis of splenic DC subsets comparing Ifnarflox/flox (WT) and CD11c Cre x Ifnar1flox/flox mice 36hrs p.i. (MVA OVA i.v.). (D, E) Representative histograms and quantitative analysis of splenic DC subsets comparing DTX-treated WT and Clec4c+/DTR animals (36hrs MVA OVA i.v.). (F, G) Proliferation profile and quantitative analysis of OT-I T cells in pLN 3d post transfer into infected (60hrs MVA OVA f.p.), DTX-treated WT, Clec4c+/DTR and Ccr5−/− animals (all groups SiglechGFP/GFP). Data are representative of at least three independent experiments (n≥3). Data are shown as mean ± standard deviation. *** = p≤0.001, ** = p≤0.01, * = p≤0.05, ns = non-significant. Please see also Figure S6.

Activated CD8 T cells recruit XCR1+ DC via XCL1

To better understand where and when pDC provided XCR1+ DC with IFN I-dependent stimulation, we analyzed the migration and localization of XCR1+ DC after infection with MVA. In the steady state, XCR1+ DC were dispersed throughout the paracortex and the interfollicular area of the LN, with some XCR1+ DC located within the subcapsular sinus (Kitano et al., 2016) (Figure 7A). Importantly, after infection we found that XCR1+ DC accumulated around OT-I T cell clusters in a pattern similar to that of recruited pDC, creating a new microenvironment around the initial site of CD8+ T cell engagement with antigen-presenting cDC (Figure 7B, 7C, 7F, 7G and Movie S7). Upon activation XCL1 is rapidly expressed by CD8+ T cells (Dorner et al., 2002), therefore we speculated that this chemokine could be critically involved in the reorganization of the XCR1+ DC network observed after infection. To address this, we transferred OT-I cells into XCR1venus/venus mice that lacked functional XCR1 expression, infected them with MVA OVA and analyzed LN sections 10hrs later. As anticipated, we did not observe a substantial cluster formation of XCR1+ DC in XCR1-deficient animals (Figure 7D, 7F, 7G). To test whether CD8+ T cell derived XCL1 is critical for XCR1+ DC recruitment, we transferred Xcl1−/− OT-I T cells into XCR+/venus mice and analyzed the XCR1+ DC localization 10hrs after infection. We observed a significant reduction in the spatial correlation between XCR1+ DC and clustered Xcl1−/− OT-I T cells as compared to WT OT-I T cells (Figure 7E–G). To further investigate the migrational behavior of XCR1+ DC, we visualized this cell population in vivo shortly after T cell cluster formation. Using this approach, we found in vivo evidence for direct migration of XCR1+ DC towards clustered OT-I T cells supporting the notion of chemokine based attraction of XCR1+ DC (Figure 7H and Movie S8).

Figure 7. Activated CD8+ T cells recruit XCR1+ DC via XCL1.

Figure 7

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(A–E) IF images showing OT-I T cell and XCR1+ DC localization in (A) the steady state, or (B–E) 10hrs p.i. (MVA OVA) in (B, C) WT (XCR1+/venus) and (D) XCR1-deficient animals (XCR1venus/venus) or (E) using Xcl1−/− OT-I T cells. (F) IF images showing OT-I T cell cluster and XCL1 staining 10hrs p.i. (G) Analysis of 3D spatial correlation between OT-I T cells and XCR1+ DC. (H) IVM images of XCR1+/venus mice showing directed XCR1+ DC migration towards an OT-I T cell cluster. Scale bars: 250μm (B), 150μm (D, E), 100μm (A), 20μm (C, H), 10μm (F). Data are representative of at least three independent experiments (n=3). Data are shown as mean ± standard deviation. *** = p≤0.001. Please see also Movie S7 and S8.

In summary, we conclude that XCR1+ DC are recruited to CD8+ T cell priming sites allowing them to interact with pDC but also with infected cDC that are localized in the middle of the cluster of T cells. This positioning and exposure to IFN I provides XCR1+ DC with a microenvironment that provides local access to cell-associated viral proteins once the initially infected DC undergoes cell death, thereby optimizing cross-presentation, maturation and productive second phase interactions with already activated CD8+ T cells.

Discussion

In this study, we analyzed the spatiotemporal dynamics of pDC to address when and where pDC-derived IFN I supports innate and adaptive immunity during viral infection. We found that pDC are directed towards two different sites in the LN as a consequence of selective and possibly competing chemokine receptor engagement. In particular, CXCL9 and CXCL10 interaction with CXCR3 directs pDC to subcapsular and medullary macrophages. It seems intuitive that this translocation to sites of viral infection may be essential to suppress viral replication, although we did not find direct evidence to support this notion. Whether CXCR3-mediated pDC migration serves another function or indeed supports the antiviral function of pDC, but was masked in our experimental setting due to an increase in pDC density within the LN, needs to be further tested.

The second area of pDC accumulation involves the priming sites of CD8+ T cells in the LN, which is the focus of this study. The critical chemokines are CCL3 and CCL4 that are produced in the context of antigen-dependent CD8+ T cell activation and engage CCR5 to direct pDC towards such initial sites of CD8+ T cell priming. The resulting accumulation of pDC around activated CD8+ T cells is likely a combination of chemotactic and adhesive cues. In this study, we have provided evidence for direct interaction between pDC and activated CD8+ T cells, which does not require cognate antigen-presentation by pDC. Similarly, pDC may interact with the infected cDC that activated naïve CD8+ T cells initially. This may be critical for pDC to sense viral infections and lead to release of IFN I, given that we did not find any evidence for direct infection of pDC. Besides pDC, we also detected a rapid recruitment of XCR1+ DC towards the activated XCL1-producing CD8+ T cells, creating a new microenvironment around the initial site of CD8+ T cell activation. Within this microenvironment, pDC-derived IFN I can act on CD8+ T cells as a signal 3 cytokines and perhaps more importantly on XCR1+ DC to optimize their maturation, costimulatory capacity, and ability to cross-present viral antigens. This in turn contributes to maximizing the fraction of the antigen-specific CD8+ T cell repertoire recruited into the immune response and/or the extent of proliferation of that cohort of CD8+ T cells encountering TCR ligand.

Taking these findings together, we propose a concept in which T cells that could locate rare antigen-bearing DC early during an infection become the nucleus of a new microenvironment that is actively created by signals emitted in response to the initial CD8+ T cell – APC interaction. Thus, rather than having to continue to search for the cells necessary to optimize their response, this mechanism permits each CD8+ T cell that locates a useful antigen-presenting cell to accumulate needed accessory cells at this existing site of activation, supporting a robust response without the risk of missing the right partner cells during further T cell migration within the lymphoid tissue.

XCR1+ DC, beyond their critical function to cross-present viral antigens, also serve as an essential platform to transmit CD4+ T cell-dependent signals to CD8+ T cells (Eickhoff et al., 2015; Hor et al., 2015). Chemokines, particularly CCL3 and CCL4, optimize cellular encounters between cDC, CD4+, and CD8+ T cells (Castellino et al., 2006; Hugues et al., 2007). In our previous work we have shown that CD4+ T cell – cDC interactions lead to CCL3 and CCL4 production, which is critical to recruitment of activated CD8+ T cells for the delivery of helper signals (Castellino et al., 2006). Given our data, the likely cellular source of CCL3 in those prior experiments was cDC rather than CD4+ T cells. Several lines of evidence argue that activated rather than naïve CD4+ and CD8+ T cells are exchanging helper signals (Eickhoff et al., 2015; Jusforgues-Saklani et al., 2008). Activated T cells typically have short-term rather than long-term interactions with cDC (Mempel et al., 2004; Miller et al., 2004; Stoll et al., 2002). Therefore, it makes sense to induce chemokine production by licensed DC to optimize CD8+ T cell recruitment for the delivery of help rather than having the source of these chemokines be CD4+ T cells that may have disengaged the licensed DC before CD8+ T cell recruitment. This is in line with a consecutive rather than simultaneous interaction model for the delivery of helper signals for CD8+ T cells (Ridge et al., 1998).

An important aspect of our study is the unraveling of how and where pDC cooperate with cDC to support adaptive immunity. Our data indicate pDC – XCR1+ DC cooperativity early after viral infection within T cell priming hubs. This is in line with previous studies demonstrating that pDC only provide IFN I early after viral infection (Swiecki et al., 2010). In our study, pDC do not seem to present antigen, but do promote cross-presentation by colocalized XCR1+ DC. Additionally, they optimize the maturation of these DC subset as shown by an upregulation of CD86, which is the critical costimulatory molecule for priming CD8+ T cells in the context of vaccinia virus infections (Salek-Ardakani et al., 2009). It is important to point out that while in our model IFN I is critical, pDC might also support adaptive immune responses through the production of other mediators such as IL-12 (Asselin-Paturel et al., 2001).

The importance of pDC and XCR1+ DC relocalization in carrying out their specific functions as shown here provides strong evidence that the capacity of pDC and XCR1+ DC to rapidly migrate within lymphoid tissues is a key aspect of their physiological function. Our data further support a concept in which in the initial CD8+ T cell – APC interaction is critical for the reorganization of the DC network, allowing for optimal cellular interactions and cooperativity to maximize CD8+ T cell priming. With our information about the spatiotemporal behavior of pDC and XCR1+ DC in LN during viral infections and the dissection of the molecular signals guiding such migration, we provide a framework for considering how to harness these cell populations to maximize cellular immunity in the context of vaccination.

Experimental Procedures

Animals

Mice were purchased from Jackson or Janvier Labs or maintained at in-house facilities. All mice were maintained in specific pathogen-free conditions at an Association for Assessment and Accreditation of Laboratory Animal Care-accredited animal facility. All procedures were approved by the NIAID Animal Care and Use Committee (National Institutes of Health, Bethesda, MD) and the North Rhine-Westphalia State Environment Agency (LUA NRW), respectively. For details on mouse strains see supplementary information.

Treatment of mice

For depletion of pDC, transgenic mice and control littermates were treated with 0.1 μg DTX i.p. (Merck Millipore) on d-2, d-1, d0, d1, d2, d3 (for analysis at d8), on d-2, d-1, d0 (for analysis 8h p.i.) and on d-2, d-1, d0, d1 i.p. (for analysis 36h p.i.), respectively. For depletion of XCR1+ DC, transgenic mice and control littermates were treated with 0.5 μg DTX i.p. (Merck Millipore) on d-1, d0 and d1. For chemokine neutralization, chemokine blocking antibodies or isotype matched controls were injected i.v. at the time of infection (R&D; CCL3: AF-450-NA, MAB450-100; CCL4: AF-451-NA, MAB451-100; Isotype matched controls: AB-108-C, MAB006) (Castellino et al., 2006). For immunization with LPS-free OVA (Hyglos), 50 μg were injected into the footpad, pLN were harvested 8h after injection.

Viral Infections

105 PFU VSV OVA or 106-108 IU MVA OVA GFP, MVA OVA tdTomato, MVA OVA or MVA WT were diluted in PBS and injected in the footpad (foothock (Kamala, 2007)), or intravenously.

Adoptive T cell transfer

OT-I, SMARTA or polyclonal CD8+ T cells were sorted using a MACS CD4+ or CD8+ T cell negative selection kit (Miltenyi) combined with biotinylated anti-CD44 (IM7, BD Biosciences). 2–4x106 cells were transferred i.v.

In vivo proliferation assay

OT-I GFP or OT-I tdTomato T cells were MACS sorted (CD8 negative selection kit, Miltenyi), labelled with Cell Proliferation Dye eFluor® 670 (eBioscience) and transferred into recipient mice 60hrs after MVA OVA infection. 72hrs post transfer pLN were harvested and single cell suspensions were analyzed by FACS.

ELISA and Chemokine Array

ELISAs and Chemokine Array of whole LN homogenates were performed according to manufacturer’s instructions (R&D Systems and pbl Assay Science).

Preparation of peptide pulsed DC

For transfer of peptide pulsed DC, splenic DC from CD11c-YFP mice were harvested, digested with collagenase and DNAse for 30min and enriched using a CD11c positive seletion kit (Miltenyi). DC were incubated in the presence of SIINFEKL peptide (μg/ml) and 5 pg/ml LPS at 37°C for 1h. After incubat ion, 6.25× 105 DC were injected into footpad. OT-I T cells were transferred i.v. 18h after injection of peptide-pulsed DC. 10h after T cell injection pLN were harvested and prepared for immunohistological analyses.

Flow Cytometry

For analysis LN and spleens were harvested and single cell suspensions were generated. For details on antibodies see supplementary information.

Immunofluorescence Staining

PLP-fixed, frozen tissues were were cut, stained, mounted and acquired on a 710 confocal microscope (Carl Zeiss Microimaging). For details on antibodies see supplementary information.

Intravital two-photon imaging

Mice were anesthetized, popliteal LNs were exposed, and intravital microscopy was performed using a protocol modified from a previous report (Kastenmuller et al., 2013a). Raw imaging data were processed using a semi-automated approach (Imaris/Bitplane). For details see supplementary information.

Statistical Analysis

Student t test (two-tailed) was used for the statistical analysis of differences between two groups with normal distribution.

Supplementary Material

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Acknowledgments

We would like to thank S. Ebbinghaus and S. Rathmann for technical assistance and D.H. Busch for kindly providing MHCI multimers. This research was supported by the Intramural Research Program, NIAID, NIH. W.Ka., N.G., C.K and W.Ko. are members of the DFG Excellence Cluster ImmunoSensation in Bonn, Germany and are supported by grant SFB704. S.D., S.B. and W.Ka. are supported by the DFG Graduate program 2168/1 (‘Bo&MeRang’). W.Ka. is supported by NRW-Rückkehrerprogramm of the German state of Northrhine-Westfalia. T.K. was supported by the Kishimoto Foundation and a Grant-in-Aid for Scientific Research (B) from Japan Society for the Promotion of Science (JSPS). F.K. is supported by the Einstein Foundation Berlin.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Author contributions: A.B., S.E. and S.D. planned and performed experiments, analyzed and interpreted the data. N.G. and W.Ko. analyzed data and designed experiments. T.K., C.K., R.K., T.Q., M.I., S.B., W.B. and M.C. were involved in study design and provided critical reagents. F.K. analyzed imaging data. R.N.G. and W.Ka. conceptualized the study, and analyzed the data. R.N.G., W-Ka. and A.B. wrote the manuscript.

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