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. Author manuscript; available in PMC: 2017 Oct 18.
Published in final edited form as: Immunity. 2016 Sep 27;45(4):847–860. doi: 10.1016/j.immuni.2016.08.019

Optimal generation of tissue-resident but not circulating memory T cells during viral infection requires crosspriming by DNGR-1+ dendritic cells

Salvador Iborra 1,*, María Martínez-López 1, Sofía C Khouili 1, Michel Enamorado 1, Francisco J Cueto 1,2, Ruth Conde-Garrosa 1, Carlos del Fresno 1, David Sancho 1,*
PMCID: PMC5074364  EMSID: EMS70264  PMID: 27692611

Summary

Despite the crucial role of tissue-resident memory T (Trm) cells in protective immunity, their priming remains poorly understood. Here, we have shown differential priming requirements for Trm versus circulating memory CD8+ T cells. In vaccinia cutaneous-infected mice, DNGR-1-mediated crosspresentation was required for optimal Trm cell priming but not for their skin differentiation or for circulating memory T cell generation. DNGR-1+ dendritic cells (DCs) promoted T-bet transcription factor induction and retention of CD8+ T cells in the lymph nodes (LNs). Inhibition of LN egress enhanced Trm cell generation, whereas genetic or antibody blockade of DNGR-1 or specific signals provided during priming by DNGR-1+ DCs, such as interleukin-12 (IL-12), IL-15 or CD24, impaired Trm cell priming. DNGR-1 also regulated Trm cell generation during influenza infection. Moreover, protective immunity depended on optimal Trm cell induction by DNGR-1+ DCs. Our results reveal specific priming requirements for CD8+ Trm cells during viral infection and vaccination.

Introduction

During infection, naive T cells in secondary lymphoid organs are primed by dendritic cells (DCs) to become either long-lived memory T cells or short-lived effectors. Based on their trafficking properties, memory T cells are subdivided among circulating memory cells and tissue-resident memory T (Trm) cells (Mueller and Mackay, 2016). Trm cells are found in all tissues and, depending on location, are characterized by stable expression of CD69, variable CD103 expression, and enhanced effector ability (Ariotti et al., 2014; Bergsbaken and Bevan, 2015; Mackay et al., 2013; Masopust et al., 2006; Schenkel et al., 2014; Skon et al., 2013; Steinert et al., 2015). Trm cells are crucial for surveying and mounting an effective and rapid immune response upon reinfection in skin and mucosae (Gebhardt et al., 2009; Jiang et al., 2012; Mackay et al., 2015; Steinert et al., 2015). Moreover, upon antigen contact, activated Trm cells orchestrate circulating memory T cell response, drive maturation of DCs, and boost local immunity, thus establishing a tissue-wide pathogen alert state (Ariotti et al., 2014; Schenkel et al., 2014; Schenkel et al., 2013).

Trm cells derive from precursors characterized by low expression of KLRG1 transcription factor that migrate into the tissue where are retained following downregulation of Krüppel-like factor 2 (KLF2) and its target gene sphingosine 1-phosphate receptor S1P1 (S1P)(Gebhardt et al., 2009; Mackay et al., 2013; Masopust et al., 2010; Skon et al., 2013), along with downregulation of T-box transcription factors (Eomes and T-bet) (Mackay et al., 2015). Trm and circulating memory T cells have the same clonal origin (Gaide et al., 2015), but Trm cells retain high-affinity T cell receptors (TCRs) (Frost et al., 2015). Whether these Trm cell precursors have specific priming requirements different from those of circulating memory cells is not known.

Cutaneous infection with vaccinia virus (VACV) in mice generates circulating memory and Trm CD8+ T cells, the latter accumulating in the skin (Jiang et al., 2012). Similarly to many viruses, VACV infects DCs and thereby provides antigens for direct presentation via MHC class I molecules (Xu et al., 2010). However, DCs can also acquire VACV antigens from infected cells, leading to CTL “crosspriming”, which contributes to prolonged antigen availability (Heipertz et al., 2014). Crosspriming to VACV largely depends on DNGR-1 (CLEC9A) (Iborra et al., 2012), a C-type lectin receptor that favors DC crosspresentation of dead-cell associated antigens (Sancho et al., 2009). DNGR-1 is highly expressed by mouse CD8α+ DCs in lymphoid organs and CD103+ DCs in non-lymphoid tissues, as well as their human equivalents (Huysamen et al., 2008; Poulin et al., 2010; Sancho et al., 2008). The development and function of CD8α+ family DCs require the action of the transcription factor Batf3 (Hildner et al., 2008; Seillet et al., 2013). Here, we report that upon VACV skin infection, Batf3-dependent DCs, operationally acting through DNGR-1-mediated crosspresentation and specific signals including IL-12, IL-15 and costimulation through CD24, are required for optimal generation of skin Trm but not effector T cells or circulating memory T cells. Moreover, DNGR-1-mediated crosspriming also controls lung Trm cell generation following influenza A virus infection, and supports effective mucosal vaccination. These data reveal priming requirements that selectively impact on the Trm cell compartment.

Results

Crosspresentation modulates CD8+ T cell-priming without affecting effector and circulating memory CD8+ T cell response

To test whether crosspresentation can affect immunity to viruses whose antigens can be both directly presented and crosspresented, we analyzed CD8+ T cell immunity to VACV intradermal (i.d.) infection comparing WT mice with two independent genetic mouse models with deficient crosspresentation of VACV antigens: DNGR-1-deficient mice (Clec9agfp/gfp) (Iborra et al., 2012) and Batf3-/- mice (Hildner et al., 2008; Seillet et al., 2013). Antigen presentation by CD11c+ DCs purified from dLN of rVACV-OVA skin-infected mice at day 1, 2 and 3 post infection (p.i.) resulted in IFN-γ production by OVA-specific OT-I CD8+ T cells (Figure 1A) or VACV-B8R-specific CTLs (Figure S1A). VACV antigen presentation was transient in WT mice (Norbury et al., 2002) and both the intensity and the persistence of antigen presentation were lower in the absence of DNGR-1 or Batf3 (Figures 1A and S1A). Next, we analyzed if defective crosspriming affects the expression of transcription factors involved in memory or effector differentiation. We found impaired induction of T-bet and its target CXCR3 during priming of OT-I T cells adoptively transferred in Batf3- or DNGR-1-deficient mice infected in the skin with rVACV-OVA (Figures 1B, 1C and S1B). Upon T cell priming, forkhead box transcription factor Foxo1 is rapidly phosphorylated and degraded (Rao et al., 2012), and we found that crosspriming deficiency resulted in weaker phosphorylation and degradation of Foxo1 (Figures 1D, 1E and S1B). In contrast, the expression of the transcription factor Eomes (Figures S1B and S1C), which promotes central memory T cell generation, were not affected.

Figure 1. Crosspresentation modulates CD8+ T cell-priming without affecting effector and circulating memory responses.

Figure 1

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(A) CD11c+ DCs were purified from dLN at the indicated time points after skin infection with rVACV-OVA (1x106 pfu) in the indicated mice. DCs were co-cultured for 6h with in vitro expanded OT-I T cells and cells were stained for CD8 and intracellular IFN-γ. Upper panel: Representative plots at 24h of co-culture in the DC:OT-I 10:1 ratio. Lower panel: Frequencies of IFN-γ+ OT-I T cells in the CD8+ compartment found at the indicated ratios DC: OT-I co-culture and time points p.i. of extraction of CD11c+ cells from dLN. Arithmetic mean ± SEM is shown, n=3 pooled experiments. (B-E) Mice of the genotype in the legend were infected with rVACV-OVA (5x104 pfu) and transferred with 105 CD45.1+ OT-I T cells. At the indicated time points T-bet (B), CXCR3 (C), phosphorylated Foxo1 (D) and Foxo1 (E) MFI was measured in OT-I T cells from the dLN by flow cytometry. (F-J) The indicated mice were infected with rVACV-OVA and transferred with OT-I T cells as in B. Frequencies (F, H) and numbers (G, I) of CD45.1+ OT-I T cells were determined 3d p.i. in the dLN in the CD8+ compartment (F, G) or in the spleen and inguinal LN (iLN) 30d p.i. (H, I). Representative plots are shown in F and H. (J) Frequencies of CD44+ CD62L+ in OTI cells in the iLN and spleen 30d p.i. (K) Mice were infected i.d. in the ear with VACV WR and draining LN cells restimulated with VACV B8R peptide or with DCs pretreated with RAW 264.7 cells infected with VACV. Frequencies of IFN-γ producing cells in the CD8+ compartment shown as individual data and arithmetic mean from a representative experiment of two performed. (L-N) The indicated mice were inoculated with MVA or not (PBS) in both ears and i.p challenged. with VACV WR 8 days later. Representative plots (L) and frequencies in the CD8+ compartment (M) of IFN-γ production by spleen cells stimulated ex vivo with B8R VACV peptide 5 days after challenge infection. (N) Frequencies of Kb-B8R20-27 VACV-specific T cells in the CD8+ compartment in the ovary of infected mice. (O-P) The indicated mice were infected with VACV WR or not (PBS) in both ears and challenged i.p. with VACV WR 30 days later. (O) Frequencies of IFN-γ production in the CD8+ compartment of spleen cells stimulated ex vivo with B8R VACV peptide. (P) Viral load in the ovary of infected mice. (F-J, M-P) Average and individual data pooled from at least two representative independent experiments and arithmetic mean are shown. * p < 0.05; ** p< 0.01; *** p < 0.001 (one way ANOVA with Bonferroni post-hoc test). See also Figure S1.

Despite impaired crosspriming, OT-I T cells transferred to mice infected in the skin with rVACV-OVA expanded normally (Figures 1F and 1G). Similarly, the frequency of endogenous circulating effector CD8+ T cells against the immunodominant VACV epitope, tracked with H-2Kb-B8R tetramers 7 d p.i. with VACV WR i.d. was comparable (Figures S1H and S1I). Next, we explored whether impaired crosspriming affects memory CD8+ T cell generation. Upon rVACV-OVA skin infection, frequency and numbers of transferred OT-I T cells, including both central and effector memory phenotype, were similar in the absence of crosspresentation (Figures 1H-J). Similarly, infection with VACV WR (i.d.) elicited comparable circulating B8R-specific CD8+ T cell memory responses (Figures S1J-S1L).

To assess the function of circulating effector and memory CTLs, we focused on DNGR-1-deficient and WT mice, since Batf3-dependent DCs play a role in CTL recall responses (Alexandre et al., 2016). Draining LN effector T cells showed similar IFN-γ production upon restimulation with VACV antigens or B8R peptide in WT and DNGR-1-deficient mice primed i.d. with VACV WR at day 7 p.i. (Figure 1K). To test whether the equivalent effector response resulted in similar viral clearance, we primed WT and DNGR-1-deficient mice with non-replicative MVA (i.d.) and subsequently infected i.p. with VACV WR 5d later. Effector responses at day 8 p.i. with VACV WR were equal in the spleen (Figures 1L and 1M) and the ovary (Figure 1N) of both mouse genotypes, which led to complete clearance of virus in the ovaries (not shown). In addition, DNGR-1 deficiency did not affect secondary responses by circulating memory CD8+ T cells analyzed in mice inoculated with VACV in the ear and i.p challenged 30 days later with the same virus. The frequency of IFN-γ+ B8R-specific CD8+ T cells in the spleen of vaccinated DNGR-1 deficient mice 5d after secondary infection (Figure 1O), as well as their viral load in the ovary (Figure 1P), was comparable to WT mice. We conclude that crosspriming increases T-bet during priming, but it is not essential for the generation of functional effector and circulating memory CD8+ T cells.

Defective crosspresentation impairs skin Trm cell responses to i.d. VACV infection

Whereas crosspresentation is dispensable for the generation of a circulating memory CD8+ T cell pool, the frequency of skin-resident (CD8+CD69+) Trm cells in the CD45+ compartment was markedly reduced in the infection site or in a distant site in the absence of DNGR-1 or Batf3 upon skin infection with VACV WR (Figure 2A). A vast percentage of these skin CD8+ Trm cells were specific for the virus and produced IFN-γ upon VACV-specific restimulation (Figure S2A). Consequently, numbers and frequencies of B8R-specific Trm cells were reduced in the skin CD45+ compartment of mice with defective crosspresentation (Figures 2B and S2B). Similarly, crosspriming-deficient mice transferred with OT-I T cells showed impaired frequency (Figure 2C and S2C) and numbers (Figure S2D) of skin Trm cells, further confirmed by immunofluorescence of skin samples 30d p.i. (Figure 2D). We analyzed the kinetics of Trm cell differentiation in the skin using CD103 as specific marker. The formation of CD103+ OT-I T cells was slower between 7 and 14 days in the absence of crosspriming, suggesting a lower number of Trm cell precursors seeded in the skin (Figure 2E).

Figure 2. Defective crosspresentation impairs skin Trm cell responses to i.d. VACV infection.

Figure 2

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(A) The indicated mice were infected with VACV WR (5x104 pfu, i.d.). Representative plots (upper panels) and individual data showing frequencies of Trm cells in the skin 30d p.i. (CD8+ CD69+ in CD45+ cells). (B) Number of Kb-B8R20-27 VACV-specific T cells in the ear 30d p.i. (C) Frequencies of OT-I Trm cells in the infected or non-infected ear 30d following transfer of 105 OT-I T cells and i.d. infection with rVACV-OVA (OT-I CD69+ in CD45+ cells). (D) Immunofluorescence staining of CD8+ T cells 30d p.i. in the infected skin of mice treated as in C. One representative image of 10 independent images acquired in two independent experiments. (E) Kinetics of CD103+ Trm cell accumulation in the skin in the indicated mice after treatment as in C. Graph shows arithmetic mean ± SEM of n=10 pooled samples from two independent experiments. (F-H) The indicated mice were infected with VACV (5x104 pfu, i.d.) or not (PBS) in the left ear, and challenged in the right ear with VACV 30 days later. Mice were treated with FTY720 at day -1, 1 and 3 after challenge infection. Representative plots (F) showing IFN-γ production by CD8+ T cells from the right ear stimulated ex vivo with B8R VACV peptide 5 days after challenge. (G) Numbers of IFN-γ+ CD8+ T cells in the ear upon restimulation with B8R peptide. (H) Viral load in the infected ear. (A-C, G, H) Arithmetic mean and individual data pooled from at least two representative independent experiments are shown. * p < 0.05; ** p< 0.01; *** p < 0.001 (one way ANOVA with Bonferroni post-hoc test). See also Figure S2.

To ask whether the net reduction in Trm cell generation in Clec9agfp/gfp or Batf3-/- mice relies on reduced priming, we mimicked it in WT mice by delaying the transfer of OT-I T cells following rVACV-OVA skin infection and infecting i.d. with VACV-WR just before OT-I T cell transfer to equalize the inflammatory environment. Delayed transfer of OT-I T cells did not affect circulating memory T cell generation (Figures S2E and S2G), while deeply impaired Trm cell generation (Figures S2F and S2H). Thus, reducing priming signals preferentially impairs Trm over circulating memory T cell generation, supporting distinct priming thresholds for generation of both compartments.

To determine whether the reduced skin Trm cell numbers found in cross-priming-deficient mice impacts their resistance to skin reinfection, we challenged WT and DNGR-1-deficient mice previously infected in one ear with a second VACV skin infection in the opposite ear 30 days after the initial infection. FTY720, a S1P inhibitor that blocks egress of circulating memory T cells from lymph nodes into blood, was administered at days -1, 1 and 3 post challenge to limit the contribution of circulating memory cells to the recall response (Jiang et al., 2012). The number of skin CD8+ T cells producing IFN-γ in response to VACV peptide (B8R-specific) was severely impaired in vaccinated DNGR-1-deficient mice compared with vaccinated WT mice, reducing Trm cell numbers to those found in non-immunized controls (Figure 2F and 2G). Impaired Trm cell generation in vaccinated DNGR-1-deficient mice resulted in defective viral clearance (Figure 2H).

DNGR-1-mediated crosspresentation is required for optimal Trm cell priming but direct presentation allows further differentiation in the skin

We explored whether the contribution of DNGR-1 and Batf3-dependent DCs to Trm cell generation was restricted to the priming step in the dLN or they could also affect Trm cell differentiation in the skin. Transferred OT-I CD8+ cells primed by skin rVACV-OVA infection in a donor mouse developed efficiently into both circulating memory CD8+ T cells and skin Trm cells only when recipient mice were also dermally infected with rVACV-OVA (Figures S3A, S3B and S3C), consistent with a recent study (Khan et al., 2016). Next, we purified primed OT-I T cells from the dLN of WT, Clec9agfp/gfp or Batf3-/- mice 3d p.i. with rVACV-OVA i.d. and transferred them to WT, Clec9agfp/gfp or Batf3-/- recipients previously infected i.d. (Figure 3A). Frequencies and numbers of circulating memory T cells generated 30d post-transfer were comparable in all cases (Figures 3A, 3B and S3D). Notably, T cells primed in WT mice gave rise to similar frequencies and numbers of Trm cells regardless of the recipient genotype, whereas T cells primed in Clec9agfp/gfp or Batf3-/- mice generated both lower frequencies and numbers of skin Trm cells in WT recipients (Figures 3A, 3C and S3E).

Figure 3. DNGR-1-mediated crosspresentation is required for optimal Trm cell priming but direct presentation allows further differentiation in the skin.

Figure 3

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(A) Experimental setup. CD45.1+ OT-I T cells were injected in WT, DNGR-1-deficient (Clec9agfp/gfp) and Batf3-/- and subsequently infected (i.d. 5x104 pfu and s.s. 1x106 pfu) with rVACV-OVA. After 72h, OT-I T cells from the dLN of WT donor mice were transferred to rVACV-OVA skin-infected WT, Clec9agfp/gfp and Batf3-/- recipients, whereas OT-I T cells from the dLN of Clec9agfp/gfp and Batf3-/- donor mice were transferred to WT mice. Representative plots showing CD45.1 and CD8 staining of inguinal LN (iLN, upper panels) and CD45.1 and CD69 in the ears (lower panels) in the recipient mice 30d after the transfer. Frequencies of (B) circulating OT-I T cells in iLN or (C) OT-I Trm cells in the CD45+ compartment in the ear. (D-E) WT mice were treated with a blocking antibody against DNGR-1 during priming (red arrows) or after priming (blue arrows). A third group of mice received a control antibody. Frequencies of Kb-8R20-27 VACV-specific T cells in the CD8+ compartment in the inguinal LN (iLN) and Trm cells in the non-infected or infected ear were assessed 30d p.i. with VACV-WR (5x104 pfu, i.d.). Graphs depict fold change of the frequencies of (D) VACV-specific circulating memory CD8+ T cells or (E) Trm cells with respect to the mean in WT mice treated with isotype control. (B-E) Arithmetic mean and individual data pooled from at least two representative independent experiments are shown. * p < 0.05; ** p< 0.01; *** p < 0.001 (one way ANOVA with Bonferroni post-hoc test). See also Figure S3.

The data above suggested that crosspriming in the LN is required for the generation of committed Trm cell precursors, but not to their differentiation in the skin. To further confirm this, we used anti-DNGR-1 blocking antibodies that block crosspriming to dead cell associated antigen in vivo (Sancho et al., 2009). As VACV antigen presentation by CD11c+ cells in the dLN takes place during the first 3d p.i. (Figures 1A and S1A and (Norbury et al., 2002)), we tested the effects of DNGR-1 blockade during or after the priming in response to skin VACV-WR infection. Blockade of DNGR-1 at any stage did not affect circulating memory T cell generation (Figure 3D). Blockade of DNGR-1 during priming, but not at later stages, impaired generation of endogenous Trm cells (Figures 3E and S3F). These results further support the important contribution of DNGR-1 for priming of Trm cell precursors using a genetic-independent approach.

Defective crosspriming leads to early T cell egress from LN

Reduced crosspriming did not influence the systemic CD8+ T cell effector response but could potentially reduce skin-homing CTLs. However, the number of effector OT-I T cells in the skin early upon rVACV-OVA skin infection was increased in Clec9agfp/gfp and Batf3-/- mice (Figures 4A and S4A). This early increase was accompanied by augmented expression of KLRG1 (Figures 4B and S4B), a marker of short-lived effector cells (Joshi et al., 2007) and increased T-bet expression in OT-I T cells in the skin of mice with defective crosspriming (Figures 4C and S4C), consistent with a negative role of high expression of T-bet during Trm cell differentiation (Laidlaw et al., 2014; Mackay et al., 2015). The predominance of short-lived effectors would explain their reduced ability to fully differentiate into CD103+ Trm cells (Figure 2E).

Figure 4. Defective crosspriming leads to early T cell egress from the LN.

Figure 4

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(A) Number of OT-I T cells in the ears of the indicated mice 4, 5 and 7d following i.d. infection with rVACV-OVA (5x104 pfu) in the ear and i.v. transfer of OT-I T cells. (B) Frequency of KLRG1+ in OT-I T cells in the ear of mice treated as in A. (C) T-bet expression (MFI) in OT-I T cells in the ear of mice treated as in A. (D) Frequency of proliferating OT-I in CD8+ T cells from peripheral blood was determined 3d p.i. in mice infected as in A and transferred with CellTrace-Violet-labeled OT-I T cells. (E, F) OT-I T cells were purified from dLN at the marked time points following i.d. infection with rVACV-OVA in the indicated mice and (E) relative expression of S1pr1 mRNA or (F) Klf2 mRNA is shown. (G, H) Mice were infected with rVACV-OVA, transferred with OT-I T cells and treated (or not) with FTY720 at day 1.5 p.i. (G) Frequencies of circulating memory OT-I T cells in the iLN 30d after i.d. infection. (H) Frequencies of OT-I CD69+ cells or (I) endogenous CD8+ CD69+ T cells in the CD45+ compartment in the ear of mice 30d after i.d. infection. Statistics is shown only for the effect of FTY720 treatment in each group. (A-D, G-I) Arithmetic mean and individual data pooled from at least two representative experiments are shown. (E, F) n=4 pooled samples of dLN OT-I T cells from 4 mice each obtained in three independent experiments. (A-H) * p < 0.05; ** p< 0.01; *** p < 0.001 (one way ANOVA with Bonferroni post-hoc test). See also Figure S4.

Increased OT-I Teff cells in the skin might indicate premature egress from the LN of newly-primed effector CTL in the absence of signals from crosspriming DCs. Indeed, we found higher frequencies of primed proliferating OT-I T cells (Figures 4D and S4D) and B8R-specific endogenous CD8+ T cells (Figure S4E) in the blood of VACV-infected Clec9agfp/gfp and Batf3-/- mice. To investigate the cause of early egress, we measured S1pr1 mRNA expression in OT-I T cells present in the dLN at different times after rVACV-OVA i.d. infection. As expected, S1pr1 expression was downregulated between 36-60h p.i. in WT mice (Figure 4E), facilitating retention of newly-primed OT-I T cells in the LN. However, S1pr1 downregulation was less persistent in the absence of crosspresentation (Figure 4E). S1pr1 expression and T cell trafficking is positively regulated by KLF2 (Skon et al., 2013). Consistent with the findings on S1pr1, the downregulation of Klf2 mRNA in OT-I T cells was more transient in crosspriming-deficient mice (Figure 4F). These data suggest that absence of crosspriming leads to early egress of CD8+ T cells.

To determine whether early egress contributes to defective Trm cell generation, we infected OT-I-transferred mice with rVACV-OVA i.d. and provided a single treatment with FTY720 36h later, thus inhibiting T cell egress to the blood (Figure S4F). The acute FTY720 treatment early p.i. increased generation of both circulating memory and Trm cells in WT mice (Figures 4G-4I and S4G-S4H). Notably, FTY720 treatment could partially rescue the defect in Trm cell generation in DNGR-1 and Batf3-deficient mice, but not to the FTY720-treated WT control (Figures 4H, 4I and S4H). These data indicate that retention of CD8+ T cells in the LN favors Trm cell generation but it is not sufficient to compensate for specific signals provided by Batf3-dependent DNGR1+ DCs.

CD8α+ and CD103+ DCs provide unique priming signals for Trm cell generation

To determine which DC subsets prime Trm cell precursors, WT mice were skin-infected with rVACV-OVA and 24h later CD11c+ cells from dLN were sorted into the CD11bhi CD8α- and CD11blo CD8α+ conventional DCs (cDCs) and CD103+ migratory DCs (mDCs) subsets. CD103+ mDCs and CD8α+ cDCs induced increased proliferation and IFN-γ production by naive OTI cells compared to CD11bhi cDCs (Figures 5A and 5B). This advantage was lost in the absence of DNGR-1, while adding exogenous pre-processed OVA peptide compensated any deficiency (Figures 5A and 5B).

Figure 5. CD8α+ and CD103+ DCs provide unique priming signals for Trm generation.

Figure 5

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(A) Representative plots showing IFN-γ staining and CellTrace-Violet dilution of naive OT-I T cells co-cultured at a ratio 10:1 DC: naive OT-I T cells for 3d with the indicated DC subsets sorted from dLN of WT mice 24h following i.d. and s.s infection with rVACV-OVA, and re-stimulated with RMA cells loaded with 1μM OVA257-264 peptide for IFN-γ production. (B) Frequencies of divided IFN-γ+ OT-I T cells in the experimental setting explained in B in which the co-culture with DCs was performed in the presence or absence of OVA257-264 peptide (5 nM) (n=3 pooled independent experiments). (C-D) Naive purified OT-I T cells were co-cultured with the indicated DC subsets from WT and DNGR-1-deficient (Clec9agfp/gfp) mice as in A. After 3d co-culture, 104 OT-I T cells were transferred to WT mice infected i.d in the ear with rVACV-OVA 3d before. Frequencies of OT-I T cells in iLN (C) or OT-I CD69+ cells in the CD45+ compartment in the ear (D) 30d after OT-I T cell transfer are shown. (E-F) Frequencies of OT-I T cells in iLN (E) or OT-I CD69+ cells in the CD45+ compartment in the ear (F) of mice treated as in D, in which OT-I:DC subset co-cultures were performed in the presence or absence of OVA257-264 (5nM). (C-F) Arithmetic mean and individual data pooled from at least two representative independent experiments are shown. (B-F) * p < 0.05; ** p< 0.01; *** p < 0.001 (B-D) one way ANOVA with Bonferroni post-hoc test (E-F) Student t test comparing OVA257-264 peptide presence or absence for each group. See also Figure S5.

To test whether in vitro priming by DNGR-1+ DC subsets was superior at generation of Trm cell precursors we transferred OT-I T cells co-cultured with DC subsets purified from rVACV-OVA infected mice as above to WT mice infected 3d before with i.d. rVACV-OVA. After 30d, transferred OT-I T cells generated comparable frequencies and numbers of circulating memory T cells regardless of the DC subset used for in vitro priming or the presence or absence of DNGR-1 (Figures 5C, S5A and S5B). Notably, CD103+ and CD8α+ DCs were superior at generating Trm cells and this occurred in a DNGR-1-dependent fashion (Figures 5D, S5C and S5D). The deficiency in the generation of Trm cell precursors by Clec9agfp/gfp CD103+ DCs was compensated by excess OVA peptide at the in-vitro co-culture stage, showing that DNGR-1-mediated crosspresentation is a key step and specifically pinpointing signal 1 provision by this DC subset as a pivotal determinant in Trm cell priming (Figures 5E, 5F and S5E). In contrast, the inability of the CD11bhi cDC subset to generate Trm cell precursors could not be rescued by excess OVA peptide (Figures 5F and S5E), supporting that DNGR-1+ DCs provide specific signals that directly prime naive CD8+ T cells to become Trm cell precursors.

IL-12, IL-15 and CD24 from DNGR-1+ DCs are required for optimal Trm but not circulating memory T cell generation

CD103+ DC and CD8α+ DCs are superior at supplying some type 2 and 3 signals that increase T-bet expression, including CD24, IL-12 and IL-15 (Kim et al., 2014; Martinez-Lopez et al., 2015; Mashayekhi et al., 2011; Muzaki et al., 2016; Sosinowski et al., 2013). In order to test their effect on Trm cell priming, we purified CD11bhi DCs and CD103+ and CD8α+ DCs from dLN of skin-infected rVACV-OVA and co-cultured them with naive OT-I T cells as above and blocking antibodies as indicated. T-bet induction in OT-I T cells was higher after co-culture with CD103+ and CD8α+ DCs compared with CD11bhi DCs, and was reduced by the blockade of IL-12, IL-15 or CD24 during the co-culture (Figures 6A and 6B).

Figure 6. IL-12, IL-15 and CD24 from DNGR-1+ DCs are required for optimal Trm but not circulating memory cell generation.

Figure 6

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(A) Representative plots showing T-bet expression of OT-I T cells co-cultured at a ratio 10:1 DC: naive OT-I T cells for 3d with the indicated DC subsets from WT mice obtained as in Figure 5A and in the presence of control antibody or blocking antibodies against IL-12, IL-15 or CD24. (B) MFI measured by flow cytometry are shown. Each dot represents one independent experiment. (C-D) Naive purified OT-I T cells were co-cultured with the indicated DC subsets, and treated with blocking antibodies as in A. After 3d co-culture, 104 OT-I T cells were transferred to WT mice infected i.d in the ear with rVACV-OVA 3d before. Frequencies of OT-I T cells in iLN (C) or or OT-I CD69+ cells in the CD45+ compartment in the ear (D) 30d after OT-I transfer are shown. (C-D) Arithmetic mean and individual data pooled from four independent experiments are shown. * p < 0.05; ** p< 0.01; *** p < 0.001 (one way ANOVA with Bonferroni post-hoc test).

The blockade of IL-12, IL-15 or CD24 did not affect the generation of circulating memory T cells 30d after the transfer of OT-I T cells primed in the co-culture with DCs (Figure 6C). In contrast, IL-12, IL-15 or CD24 blockade impaired the priming of Trm cell precursors by DNGR-1+ DCs (Figure 6D). These results support that, together with increased and prolonged signal 1 via DNGR-1, additional signals from CD103+ and CD8α+ DCs that promote T-bet expression during priming (including CD24, IL-12 or IL-15) selectively favor Trm over circulating memory T cell generation.

Deficient crosspriming selectively impairs lung Trm cell generation and vaccination

To extend our results, we tested generation of circulating memory T cells and lung Trm cells following intranasal (i.n.) infection with influenza A virus. We tracked lung CD8+ Trm cells 40d after influenza PR8 virus infection (Laidlaw et al., 2014) (Figure S6A). DNGR-1 deficiency resulted in impaired endogenous generation of Trm cells specific for two influenza epitopes analyzed (NP and PA) (Figure 7A, and S6B-S6D). In the case of NP, CD103+ and CD103- Trm cells were generated, whereas for PA, most Trm cells generated were CD103+. In contrast, DNGR-1 deficiency did not affect generation of NP-specific circulating memory T cells in the inguinal LN, while PA-specific circulating memory T cells were even increased (Figure 7B and S6E-S6G). These results further support the notion that priming of Trm cells and circulating memory T cells is differentially regulated by DNGR1+ DCs in several viral infection models.

Figure 7. Deficient crosspriming selectively impairs lung Trm cell generation and vaccination.

Figure 7

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(A-B) Memory CD8+ T cells were assessed 40 days after i.n. challenge with influenza A virus PR8 (0.2xLD50) in WT and DNGR-1-deficient (Clec9agfp/gfp) mice. (A) Representative plots and frequencies in the CD8+ compartment of CD103+ and CD103- Db-NP366-374 tetramer positive in lung-localized (unlabelled CD8β cells) CD45+CD8+ T cells (see Figure S6A). (B) Frequencies of Db-NP366-374 tetramer-positive in the circulating memory CD8+ T cells in iLN. (C-D) Mice were infected i.n. with rVACV-NP and analyzed 60d p.i. (C) Representative plots and frequencies of Db-NP366-374 tetramer-positive circulating memory CD8+ T cells in the inguinal LN (iLN). (D) Representative plots and frequencies of Db-NP366-374 tetramer positive CD8+ T cells in lung-localized CD45+cells. (E-F) Mice were immunized with 5×103 pfu of rVACV-NP or VACV-WR and 60d later challenged with influenza A virus PR8 (2xLD50). (E) Survival curves (p= log rank test) (F) weight loss, mean ±s.e.m. (A-D) Arithmetic mean and individual data pooled from at least two representative independent experiments are shown. (E-F) n=10, one experiment representative of two performed. (A-D, F). * p < 0.05; ** p< 0.01; *** p < 0.001 (A, B, F) Student’s t test; (C-D) one way ANOVA with Bonferroni post-hoc test). See also Figure S6.

Next, we investigated whether a recombinant VACV expressing the influenza nucleoprotein (rVACV-NP) could generate NP-specific Trm cells in a DNGR-1 and Batf3-dependent fashion. Following rVACV-NP intranasal (i.n.) administration, crosspresentation deficiency did not affect the early CD8+ T cell effector responses (Figure S6H) or circulating memory CD8+ T cell compartment in the LN (Figure 7C). In contrast, the lung-resident CD8+ T cell memory compartment was severely depleted in the absence of DNGR-1 or Batf3 (Figure 7D and S6I).

To test the effect of deficient Trm cell generation on vaccination, we used rVACV-NP administered i.n. as a vaccine vector against influenza A virus. In this model, protection from influenza virus infection is dependent on memory CD8+ T cells specific for NP (Slütter et al., 2013). We found that, in contrast to WT mice, Clec9agfp/gfp and Batf3-/- mice were not protected against pulmonary influenza virus challenge 60d after i.n. infection with rVACV-NP (Figure 7E and 7F). In conclusion, protective mucosal immunity depends on optimal Trm cell induction by DNGR-1+ DCs.

Discussion

The induction of long-lived cell-mediated immunity in non-lymphoid tissues is a major challenge for rational vaccine design. Recent contributions focus on how Trm cells are differentiated in non-lymphoid tissues (Bergsbaken and Bevan, 2015; Mackay et al., 2013; Mackay et al., 2015; Skon et al., 2013; Wakim et al., 2010; Wakim et al., 2012), but specific priming requirements for Trm cell generation are unknown. Fate decision between effector and memory CD8+ T cell differentiation occurs early upon T cell priming (Iborra et al., 2013; Kim et al., 2014; Rao et al., 2010). The reconstitution of mature Trm cells upon adoptive transfer with a KLRG1lo subset from day-7 spleen supports the existence of an imprinted Trm precursor generated in secondary lymphoid organs (Mackay et al., 2013). Our data support that Trm and circulating memory cell precursors are generated together in the LN, consistent with a common naive T cell precursor for both subsets (Gaide et al., 2015). Our results indicate that strength and duration of specific priming signals from DCs differentially affect these TCR-identical cells to generate circulating versus tissue-resident memory CD8+ T cells.

Crosspriming results in transient inactivation of Foxo-1 in CD8 T cells favoring their retention in the LN. Foxo1 inactivation depends on the kinase mTORC1 (Rao et al., 2012) and rapamycin, a well-known mTORC1 kinase inhibitor, enhances circulating memory T cell precursor differentiation in vivo when administered at low doses (Araki et al., 2009; Pearce et al., 2009), but selectively impairs Trm cell generation (Sowell et al., 2014). Crosspriming also induced transiently T-bet correlating with generation of Trm cell precursors in the LN. While low-affinity T cells, which exhibit low expression of T-bet during priming, preferentially differentiate into central memory cell precursors (Knudson et al., 2013), Trm cells are enriched in high affinity TCRs compared with their circulating counterparts (Frost et al., 2015). T-bet induction at priming may also favor longer retention in the LN (Mackay et al., 2015) of T cells that finally egress with lower T-bet and KLRG1 expression. Indeed, we found higher expression of T-bet and KLRG1 in skin T cells of cross-priming deficient mice, consistent with a negative role of high expression of T-bet during Trm cell differentiation in the skin (Laidlaw et al., 2014; Mackay et al., 2015). Notably, impaired crosspriming does not impact Eomes expression, which would explain the lack of an effect on circulating memory T cell generation (Iborra et al., 2013; Joshi et al., 2007; Rao et al., 2010).

Consistent with previous reports (Xu et al., 2010), our current data support that direct presentation is sufficient for generation of a fully functional circulating effector and memory response against most VACV-derived peptides. DNGR-1-mediated crosspriming is required for optimal generation of effector CTLs against peptides that are highly or strictly dependent on cross-presentation, as we previously described (Iborra et al., 2012), and to induce Trm cell precursors that generate functional Trm cells that promote resistance upon reinfection in barrier tissues. Crosspresentation may prolong antigen availability for CD8+ T cells (Heipertz et al., 2014), which has been shown to determine fate decisions during priming (Henrickson et al., 2013). Our data has shown that the strength and persistence of antigen presentation was impaired in the absence of DNGR-1 crosspriming, preferentially decreasing the generation of Trm over circulating memory T cells. These results suggest that the priming threshold for the generation of Trm cell precursors differs from that for circulating memory T cell precursors. Reduced availability of antigen and/or activating signals over time has been linked to memory differentiation, whereas increased stimulation over time promotes effector and/or effector-memory differentiation (Badovinac et al., 2007; D'Souza and Hedrick, 2006). Our results suggest that Trm cell precursors are more related to an effector differentiation pathway as they require strong priming by Batf3-dependent DCs.

The KLF2-dependent receptor S1P1 is downregulated after T cell activation, leading to retention in the dLN during priming (Carlson et al., 2006). When priming signals are decreased, KLF2 and S1P1 are upregulated, allowing primed T cells to egress from the LN and access the skin, where inflammatory signals again result in KLF2 and S1P1 downregulation (Skon et al., 2013). We found that downregulation of KLF2 and S1P1 during priming in the LN was more transient in the absence of crosspresentation. As a probable consequence, T cells exit to the blood in greater numbers and are recruited to the skin earlier, but as they are KLRG1+ they may not generate Trm cells in the skin (Mackay et al., 2013), but rather behave as terminal effectors (Joshi et al., 2007; Sarkar et al., 2008). Supporting this, we found increased skin effector T cell contraction rate and reduced rate of generation of Trm in crosspriming deficient mice. Inhibition of S1P1-mediated egress with FTY720 resulted in significantly more Trm cells in WT mice. However, inhibition of egress was not sufficient to fully rescue the impaired Trm cell generation in the absence of crosspresentation.

Our data show that priming of naive CD8+ T cells to induce Trm cell precursors was mediated by Batf3-dependent DCs in the LN and required DNGR-1. However, Batf3 and DNGR-1 were not required for local differentiation of Trm cell precursors in the skin, which also required antigen presentation. It is conceivable that different DC subsets work cooperatively to promote priming in the LN and differentiation in the skin, as human lung tissue-resident CD1c+ DCs, but not Batf3-dependent CD141+ DCs, drive differentiation of CD8+ T cells with a Trm cell phenotype (Yu et al., 2013). This need of antigen for Trm cell lodgment and differentiation in tissues has been described (Wakim et al., 2010), but is not found in other systems (Casey et al., 2012; Mackay et al., 2013; Mackay et al., 2012), suggesting that priming or differentiation requirements depend on the pathogen or inflammatory insult (Gaide et al., 2015). In fact, a remaining Trm cell fraction in our system can be still generated in the absence of cross-priming. Notably, supplementation with pre-processed antigen in vitro did not rescue the inability of CD11bhi DCs to generate cell Trm precursors, indicating that only CD103+ or CD8α+ crosspresenting DCs can provide the appropriate signals for priming Trm cells during VACV infection. There are specific signals that qualitatively distinguish priming via Batf3-dependent DCs versus CD11bhi DCs, including CD24, IL-12 and IL-15 (Kim et al., 2014; Martinez-Lopez et al., 2015; Mashayekhi et al., 2011; Muzaki et al., 2016; Sosinowski et al., 2013). We found that blockade on the priming capacity of DNGR-1+ DCs mediated by any of these signals impaired T-bet expression during priming and Trm but not circulating memory T cell generation. Our data indicate that priming in cis by DNGR-1+ DCs is needed for Trm cell generation, which occurs operationally via DNGR-1-mediated crosspriming and may depend on the nature of the infection (Desch et al., 2014).

Our results contribute to open research avenues investigating the differential factors needed for improving priming of Trm cell responses and thus more effective skin or mucosal CD8+ T cell vaccination. Optimal immunization would require adequate access of antigen and adjuvants to promote priming via Batf3-dependent DCs, conditions that enhance Trm cell generation without negatively affecting circulating memory T cell generation.

Experimental Procedures

Mice

Mouse colonies were bred at the CNIC in specific pathogen-free conditions. Clec9agfp/gfp mice (DNGR-1-deficient) (Sancho et al., 2009) and Batf3-/- mice (kindly provided by Dr. K. M. Murphy, Washington University, St. Louis, MO) (Hildner et al., 2008) were backcrossed to the C57BL/6 background. OT-I transgenic mice (C57BL/6-Tg(TcraTcrb)1100Mjb/J) were mated with B6-SJL (Ptprca Pepcb/BoyJ) expressing CD45.1 allele, both from The Jackson Laboratory. Animal studies were approved by the local ethics committee. All animal procedures conformed to EU Directive 2010/63EU and Recommendation 2007/526/EC regarding the protection of animals used for experimental and other scientific purposes, enforced in Spanish law under Real Decreto 1201/2005.

Viral infections, adoptive transfer and FTY720 treatment

VACV WR strain, rVACV expressing full length OVA and the rVACV containing the influenza A/Puerto Rico/8/34 virus nucleoprotein were a gift from J. W. Yewdell and J. R. Bennink (NIH, Bethesda, MD) and were kindly provided by M. del Val (CBMSO, Madrid). Stocks were grown and virus titration was performed as described (Iborra et al., 2012). Mice were infected i.n. (5x103 pfu), or i.d. into the ear pinnae (5x104 or 1x106 pfu) or by skin scarification (s.s.) in the base of tail (1x106 pfu) or i.p. (1x106 pfu) with the stated VACV strain or i.d. (1x105) with MVA (B. Moss, NIH, Bethesda, MD), where indicated. Influenza A/Puerto Rico/8/34 PR8 virus was kindly provided by E. Nistal-Villán (University CEU San Pablo, Madrid). Naive OT-I T cells (105), in vivo primed OT-I T cells (105), in vitro activated OT-I T cells (104) purified by negative selection or B8R-specific CD8 T cells (5x103) sorted by flow cytometry were adoptively transferred to mice where indicated. FTY720 (Cayman Chemical) was inoculated i.p. (7 mg/kg) in aqueous solution.

Processing of ears, lungs and dLN

Ears, lungs or LNs were harvested from naive mice or mice infected at the indicated times and single-cell suspensions were prepared by liberase/DNAse digestion. DCs from LN were enriched using anti-CD11c-microbeads (Miltenyi Biotec). CD8+ T cells were enriched by negative selection using a cocktail of biotin-conjugated antibodies (anti-CD11c, CD11b, B220, MHC-II, CD4, NK1.1) followed by separation with Streptavidin-microbeads (Miltenyi Biotec). T cells were restimulated to induce cytokine production by co-culture with DCs for 6h, and brefeldin A (Sigma, 5 μg/ml) added for the last 4h of culture. When needed, enriched CD11c+ cells were further sorted into CD11bloCD8α+CD103-, migratory CD11bloCD8α- CD103+and CD11bhi CD8α- CD103- DCs in a BD FACSAria Sorter or a Sony Synergy 4L Cell Sorter. Where indicated, DC subsets were co-cultured with naive OT-I T cells in the presence or not of blocking antibodies to IL-12 (C17.8; In Vivo Ready™ Tonbo Biosciences), IL-15 (AIO.3, eBioscience), CD24 (clone M1/69, gifted by C. Ardavín) or control antibody (rat IgG).

Peptides, tetramers, antibodies and flow cytometry

The 20TSYKFESV27 (B8R) peptide was a kind gift from Hisse M. Van Santen (CBMSO, Madrid). The peptide 257SIINFEKL264 from ovalbumin was purchased from GenScript. APC-labeled tetramers specific for VACV, H-2Kb (20TSYKFESV27, B8R), and for Influenza A virus, H2-Db (366ASNENMETM374, NP) and H2-Db (224SSLENFRAYV233, PA) were provided by the NIH Tetramer Facility at Emory University. APC-labeled dextramers specific for OVA H-2Kb (257SIINFEKL264) were purchased from Immudex.

Samples for flow cytometry were stained in ice-cold PBS supplemented with 2mM EDTA, 1% FCS and 0.2% sodium azide, with the appropriate antibody cocktails. Anti-mouse CD45, CD4, CD8α, CD8β, CD11b, CD11c, CD103, CD62L, CD44, CXCR3, KLRG1, Eomes, T-bet, IFN-γ and I-Ab (MHC-II) antibodies conjugated to biotin, FITC, PE, PerCP-Cy5.5, V450 or APC were obtained from eBioscience, BD Pharmingen and Biolegend. APC-Cy7 CD8 was from Tonbo Biosciences. For proliferation assays, OT-I T cells were fluorescently labelled (5 μM, CellTraceTM Violet). Antibodies to Foxo1 phosphorylated at Ser256 and total Foxo1 antibody (L27) were from Cell Signaling. Intracellular staining of transcription factors was performed using the Foxp3-fixation-permeabilization buffer from eBioscience. Events were acquired using a FACSCanto flow cytometer or FACSDiva (Becton Dickinson) and data were analyzed using FlowJo software (Tree Star). Intravascular staining was performed as described (Laidlaw et al., 2014). Briefly, a total of 3 μg anti–CD8β was injected i.v. At 3 min after injection, the animals were sacrificed, bled, and perfused with 10 ml cold PBS. The LNs and lung were harvested and lymphocytes were isolated as described.

Statistical analysis

The statistical analysis was performed using Prism software (GraphPad Software, Inc). Variance equality among groups was determined using F-test. Statistical significance for comparison between two groups of samples coming from a normal distribution (Shapiro-Wilk test for normality) was determined using the unpaired two-tailed Student's t test. For comparison of more than two groups, one way ANOVA and Bonferroni post-Hoc test was used. A p value <0.05 was considered significant. Mice were used randomly in the different experimental procedures.

Supplementary Material

Suppl. Mat.

Acknowledgements

We are grateful to C. Reis e Sousa, E. Fernández-Malavé, M. del Val, J. Boettcher, M. Robinson and members of the DS lab for discussions and critical reading of the manuscript. We thank the CNIC facilities, personnel and to S. Bartlett for editorial assistance. We acknowledge the NIH Tetramer Core Facility (contract HHSN272201300006C) for MHC-I tetramers. SI is funded by grant SAF2015-74561-JIN. Work in the DS laboratory is funded by the CNIC and grants from the Spanish Ministry of Economy and Competitiveness (MINECO, SAF-2013-42920R), and the European Research Council (ERC-2010-StG 260414). The CNIC is supported by the MINECO and the Pro-CNIC Foundation, and is a Severo Ochoa Center of Excellence (MINECO award SEV-2015-0505).

Footnotes

Author contributions

S.I., M. M-L., S.C.K, M.E., F.J.C., R.C-G. and C. del F. did the experiments; S.I. and D.S. conceived and designed experiments, analyzed data and wrote the manuscript. All the authors discussed the results and the manuscript.

The authors declare no competing financial interests.

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