Classical Type 1 Dendritic Cells Dominate Priming of Th1 Responses to Herpes Simplex Virus Type 1 Skin Infection

CD4⁺ T cells mediate protection from a broad range of pathogens. Efficient immunity against intracellular pathogens often requires CD4⁺ T cells that are capable of producing cytokines such as IFN-γ. However, the mechanisms driving the generation of CD4⁺ T cells with appropriate functional polarization remain incompletely defined, which in turn limits our ability to design T cell vaccines. Although Ag presentation by dendritic cells (DC) is pivotal to the generation of T cell responses, there are several variables involved in this process, including the nature of the Ag, how it is acquired, and the type of DC interacting with the T cells, all of which influence the nature of the ensuing immune response.

Conventional DC subsets are commonly categorized based on their anatomical location, surface marker expression, and developmental origin. In murine skin and skin-draining lymph nodes (LN), these include the epidermal Langerhans cells (LC), CD103⁺ dermal DC (dDC), CD103^– dDC, and LN-resident CD8a⁺ DC (1–3). CD103⁺ DC and CD8α⁺ DC have recently been grouped together as classical type 1 DC (cDC1) based on their numerous shared traits, including a developmental dependence on the transcription factor IFN regulatory factor 8 (IRF8) and the ability to secrete the cytokine IL-12, which promotes Th1 responses (1–10). Indeed, several reports have implicated lymphoid CD8α⁺ and migratory CD103⁺ DC as promoters of Th1 responses to pathogens, whereas other conventional DC have been shown to stimulate Th17 and Th2 responses (5, 8, 11–14).

The induction of CD4⁺ T cell proliferation and differentiation in vivo involves multiple interactions with Ag-bearing DC (15–19). Therefore, the functional status and type of the Ag-presenting DC, the nature and abundance of the Ag, and the sequence with which T cells encounter distinct DC all have the potential to impact on the polarization of the ensuing effector T cell response. Recent work has suggested that following skin infection with HSV type 1 (HSV-1) in mice, Ag is initially presented to CD4⁺ T cells by DC that have migrated from the skin, and subsequent to this, CD4⁺ T cells interact with LN-resident CD8α⁺ DC (20, 21). Although previous reports have suggested that migrating and LN-resident DC play distinct roles in driving CD4⁺ T cell expansion in LN after s.c. Ag delivery (22, 23), how such spatially and temporally separated interactions of CD4⁺ T cells with these DC impact on functional T cell polarization and cytokine production capacity after skin infection remains to be determined.

In this article, we show that inoculation of skin with an inactivated nonreplicative virus elicited impaired CD4⁺ T cell responses in part due to the presentation of viral Ag being limited to CD103⁻ dDC. Likewise, impaired Th1 responses were observed following skin infection with replicating virus in mice lacking expression of IRF8 required for the development of cDC1, including LN-resident CD8α⁺ DC. Finally, direct targeting of Ag to cDC1 shortly after inoculation with inactivated virus resulted in the restoration of Th1 cell differentiation. Collectively, the data suggest that optimal Th1 responses to localized virus infection of the skin are induced by the cooperative priming by both migratory and LN-resident DC subsets.

C57BL/6 (B6), OT-II (24), OT-II.dsRed, gDT-II×B6.CD45.1 (gDT-II) (20), Irf8^−/−(25), Il12p40^−/− (10), and Xcr1^+/venus mice (26) were used in this study. All mice were bred and/or maintained in the Department of Microbiology and Immunology of the University of Melbourne, Parkville. All animal experimentation was approved and complied with The University of Melbourne Animal Ethics committee regulations. All mice were female unless otherwise stated in the figure legend.

gDT-II and OT-II cells were isolated from pooled LN and spleens from donor mice. Organs were extruded through a 70-μm nylon mesh to generate a single-cell suspension. RBCs were removed using RBC lysis buffer (Sigma-Aldrich). CD4⁺ T cells were enriched using a depletion mixture that contained rat Abs to Mac-1 (M1/70), F4/80 (F4/80), erythrocytes (TER-119), I-A/E (M5/114), and CD8α (53.6-7), with Ab-bound cells removed using goat anti-rat IgG-coupled magnetic beads (Qiagen). Further positive enrichment of CD4⁺ T cells was performed by incubating cells in CD4 enrichment Dynabeads mix (Invitrogen), followed by DETACHaBEAD (Invitrogen) treatment to remove magnetic beads. The purity of enriched cells was ∼50% for gDT-II cells (identified by cell-surface expression of CD4 and Vα3.2 TCR) and >90% for OT-II cells (identified by cell-surface expression of CD4 and Vα2 TCR). Cells were enumerated and adjusted to 5 × 10⁵ cells (proliferation assays and the cytokine production assay comparing HSV-1 and UV-HSV inoculation of wild-type mice), 1–2 × 10⁵ cells (confocal microscopy), or 1 × 10⁴ cells (all other in vivo assays) for adoptive transfer into mice at least 24 h prior to inoculation with virus. B cells from OT-II.dsRed donor mice were obtained by depletion of CD43⁺ splenocytes using anti-CD43 magnetic microbeads (MACS; Miltenyi Biotec). To readily denote B cell areas in LN for confocal microscopy analysis, 2 × 10⁶ dsRed B cells were transferred into recipient mice together with enriched gDT-II and OT-II cells.

Enriched gDT-II and OT-II cells were incubated with 5 μM CellTrace Violet (CTV; Invitrogen) or 500 nM of 1:1 CellTracker Deep Red (CTDR)/10% Pluronic F-127 (Invitrogen), respectively, in PBS solution with 0.1% (v/v) heat-inactivated FCS at 37°C for 10 min. The reaction was quenched by the addition of ice-cold FCS, and cells were extensively washed with and resuspended in either HBSS alone for injection into mice via the tail vein or RPMI 1640 medium supplemented with 10% (v/v) FCS, 2-ME (50 μM), ʟ-glutamine (2 mM), penicillin (100 U/ml), and streptomycin (100 μg/ml) for culture with sorted DC subsets.

HSV-1 KOS and SC16 strains were grown in MEM with 10% (v/v) FCS. HSV-1 was diluted to working concentration prior to inactivation by 30 min exposure to UV light 10 cm from the source. Epicutaneous skin inoculation with either live HSV-1 or UV-HSV involved administration of 1 × 10⁶ PFU to abraded flank skin as previously described (27). Infectious virus in the tissue, virus stock titration, and inactivation were measured by standard assay of PFU on Vero cells (CSL) as previously described (27). HSV-1 KOS strain was used for all experiments unless otherwise stated.

Single-cell suspensions were stained in PBS with 2% FCS (v/v) and 5 mM EDTA in the presence of the Fc receptor blocking Ab (2.4G2) with Abs specific for the following Ags: CD4 (GK1.5), CD45.1 (A20), Vα3.2 (RR3-16), CD44 (IM7), CD25 (PC61.5; eBiosciences), CXCR3 (220803; R&D Systems), T-box expressed in T cells (T-bet; eBio4B10; eBioscience), IFN-γ (XMG1.2), TNF-α (MP6-XT22), and IL-2 (JES6-5H4). Abs were obtained from BD Biosciences unless otherwise stated. For T-bet detection, cells were stained using a Foxp3 staining buffer kit (eBioscience). For detection of cytokines, cells were fixed with 1–2% paraformaldehyde (Electron Microscopy Sciences) and permeabilized in PBS with 2% FCS (v/v) and 5 mM EDTA supplemented with 0.4% saponin (Sigma-Aldrich). All flow cytometry experiments were performed on LSRFortessa, LSR II, or FACSCanto II flow cytometers (BD Biosciences). Data were analyzed using FlowJo (Tree Star). Dead cells were excluded from the analysis by staining cells with propidium iodide (Sigma-Aldrich), 7-AAD (Sigma-Aldrich), or Fixable Viability Dye (eBioscience).

Single-cell suspensions from brachial LN (bLN) of inoculated mice at 2 or 3 d postinoculation (dpi) were produced by digestion in RPMI 1640 medium supplemented with 2.5% (v/v) FCS containing 1 mg/ml collagenase type III (Worthington) and 0.02 mg/ml DNase I (Roche). LN suspensions were enriched for DC with the following Abs: anti-Gr1 (RB6-8C5), anti-CD3ε (KT3), anti-CD19 (ID6), anti-Thy1 (T24), anti-B220 (RA36B2), and anti-Ly76 (Ter-119), followed by removal using goat anti-rat IgG-coupled magnetic beads (Qiagen). Cells were stained with anti-CD11c (N418; BD Biosciences), anti-CD205 (205yekta; eBioscience), anti-CD326 (G8.8; BioLegend), anti-CD103 (M290; BD Biosciences), and anti-CD8α (53-6.7; BD Biosciences) and sorted into CD8α⁺ DC, CD103⁺ dDC, LC, and CD103⁻ dDC. After sorting, DC subsets were washed and resuspended in RPMI 1640 medium supplemented with 10% (v/v) FCS, 2-ME (50 μM), ʟ-glutamine (2 mM), penicillin (100 U/ml), and streptomycin (100 μg/ml) and cultured together with gDT-II cells as previously described (20, 28). A similar DC enrichment method was employed for flow cytometry analysis of DC subsets in B6 and Irf8^−/− bLN; however, cells were additionally stained with anti-CD45.2 (104; BD Biosciences) and anti–MHC class II (MHC II) (M5/114.15.2; eBioscience) for these experiments.

Cells were cultured at 37°C in DMEM supplemented with 10% (v/v) FCS, 2-ME (50 μM), ʟ-glutamine (2 mM), penicillin (100 U/ml), and streptomycin (100 μg/ml) for 5 h total in the presence or absence of 5 μM gD_315–327 peptide (GenScript). After 1 h, 10 μg/ml Brefeldin A (Sigma-Aldrich) was added to all samples.

Two micrograms of recombinant glycoprotein D from HSV-1 fused to either the H chain of rat anti-mouse Clec9A (clone: 24/04-10B4) or to a rat IgG2a isotype control (clone: GL117) in PBS was i.v. injected via the tail vein as previously described (29).

bLN were harvested 2 dpi from Xcr1^+/venus mice that had been adoptively transferred with gDT-II (labeled with CTV), OT-II (labeled with CTDR), and B cells (dsRed) and fixed in paraformaldehyde-lysine-periodate fixative for 2–6 h prior to embedment in 2% agarose. Thick 200–250 mm LN sections were produced using a VT1200 S vibratome (Leica Biosystems). Three-dimensional tiled images were acquired and automatically stitched together on an LSM710 NLO multiphoton microscope using Zen Imaging software (Carl Zeiss). Subsequent processing and analysis of the three-dimensional mosaic images were completed using Imaris (Bitplane). The Imaris spot creation module was used to determine the location of each adoptively transferred T cell, and clustering cells were defined as those that were within 20 μm of at least one other like cell.

The primary inoculation site, defined as a 1 cm² piece of skin encompassing the area of scarification, and the secondary site, which was 1 cm wide and extended from 5 mm below the primary site to the ventral midline of the mouse, were excised and homogenized in serum-free DMEM prior to performing PFU assays.

Single-cell suspensions of bLN from inoculated mice at 2 dpi were produced by digestion in RPMI 1640 medium supplemented with 2.5% (v/v) FCS containing 1 mg/ml collagenase type III (Worthington) and 0.02 mg/ml DNase I (Roche). bLN suspensions were enriched for DC with Abs to anti-Gr1 (RB6-8C5), anti-CD3ε (KT3), anti-CD19 (ID6), anti-Thy1 (T24), anti-B220 (RA36B2), and anti-Ly76 (Ter-119), followed by removal using goat anti-rat IgG-coupled magnetic beads (Qiagen). Cells were stained with anti-CD3ε (145-2C11; eBioscience), anti-CD19 (ID6; BD Biosciences), anti-NK1.1 (PK136; BD Biosciences), anti-B220 (RA36B2; eBioscience), anti-CD11c (N418; BD Biosciences), anti–MHC II (M5/114.15.2; eBioscience), anti-CD205 (205yekta; eBioscience), anti-CD326 (G8.8; BioLegend), anti-CD103 (M290; BD Biosciences), anti-CD8α (53-6.7; BD Biosciences), anti–XCR-1 (ZET; BioLegend), anti-CD11b (M1/70; BD Biosciences), anti-SIRPα (P84; BioLegend), and anti-CD40 (3/23, BD Biosciences) and were analyzed by flow cytometry.

Statistical significance was determined via Prism 7 (GraphPad), and the specific statistical tests used are indicated in the figure legends. Symbols denoting significance are ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05, and NS, p ≥ 0.05.

To investigate the cellular and pathogen-related requirements for the generation of robust antiviral CD4⁺ T cell immunity, we used a murine model of epicutaneous HSV-1 infection in which CD4⁺ T cells are critical for both optimal humoral and cellular immunity (30–32). We further wished to modify parameters such as Ag load, Ag distribution, and inflammation and assess their impact on the resulting CD4⁺ T cell response. Therefore, we began by comparing the expansion of adoptively transferred HSV-specific CD4⁺ transgenic T cells (gDT-II) following inoculation of murine flank skin with either live, infectious HSV-1, or virus inactivated by exposure to UV radiation (UV-HSV). Infectious HSV-1 stimulated the proliferation of gDT-II cells, with numbers in the draining bLN peaking at 7 dpi (Fig. 1A, 1B). In contrast, exposure to inactivated virus elicited a markedly different pattern of expansion. gDT-II cell numbers in bLN peaked at 5 dpi; however, the magnitude of the response was strongly reduced relative to infection with live HSV-1 and subsided quickly thereafter. Impaired gDT-II expansion after UV-HSV exposure further translated to strongly diminished numbers of gDT-II cells recovered from spleen (Fig. 1C). Thus, replicative HSV-1 elicited a CD4⁺ T cell response that differed in kinetics and magnitude from that elicited by inactivated virus.

Next, we assessed whether the reduced CD4⁺ T cell response to UV-HSV was coupled with altered T cell activation and differentiation. Focusing on the early response, gDT-II cells labeled with the proliferation dye, CTV, were adoptively transferred into mice that were subsequently inoculated with either HSV-1 or UV-HSV (Fig. 2A). Although no division of gDT-II cells was observed within the bLN of naive controls, gDT-II cells responding to either HSV-1 or UV-HSV had divided extensively by 3 dpi (Fig. 2B, 2C). In all cases, inoculation with UV-HSV induced fewer gDT-II cells to divide, with a significantly lower proportion of gDT-II cells undergoing more than six divisions compared with gDT-II cells responding to bona fide infection (Fig. 2C). Moreover, following infection with HSV-1, gDT-II cells increased expression of the hallmark activation and Th1 differentiation markers, CD44, CD25, CD122, and CXCR3. Interestingly, there were only marginal changes in the expression of CD25, CD122, and CXCR3 as gDT-II cells divided following exposure to UV-HSV, despite the fact that these cells had undergone multiple rounds of division and upregulated the expression of CD44 (Fig. 2D, 2E).

The expression of CD122 and CXCR3 is regulated by the Th1 master transcriptional regulator, T-bet (33–36). In line with this, we noted that T-bet expression was induced in gDT-II cells isolated 3 dpi from mice infected with HSV-1, whereas those isolated from mice that received UV-HSV had only a negligible amount of T-bet expression (Fig. 2F, 2G). Moreover, consistent with the critical role of T-bet in the production of IFN-γ (35, 36), only gDT-II cells isolated from mice infected with live HSV-1 readily produced IFN-γ upon restimulation with cognate peptide Ag (Fig. 2H, 2I). Interestingly, a considerable proportion of gDT-II cells isolated from mice that received UV-HSV produced IL-2. Nevertheless, this proportion was lower than that observed in infected mice (Fig. 2H, 2J). Taken together these data indicate that inoculation of skin with UV-HSV stimulates division of HSV-specific CD4⁺ T cells; however, this response was associated with an aberrant phenotype, characterized by reduced expression of key components of the IL-2R complex, T-bet, and IFN-γ.

Several reports have suggested that optimal CD4⁺ T cell expansion and differentiation requires prolonged Ag presentation and several successive interactions with DC during the inductive phase of the response (15–17, 19, 22). Therefore, we hypothesized that the impaired expansion and aberrant differentiation of gDT-II cells induced by UV-HSV may have resulted from a waning or even a loss of Ag presentation during the 3 d assay period following exposure to UV-HSV. To test if inoculation with UV-HSV resulted in curtailed Ag presentation, mice received either HSV-1 or UV-HSV followed by CTV-labeled gDT-II cells i.v. at 3, 5, or 9 dpi, and the in vivo proliferation of the gDT-II cells was assessed 50 h later (Fig. 3A). Proliferation of gDT-II cells was evident when cells were transferred 3 dpi irrespective of whether the mice received HSV-1 or UV-HSV, although exposure to replicative HSV was associated with an increased proportion of cells in later divisions and significantly higher numbers of divided gDT-II cells (Fig. 3B–D). These differences became more pronounced at later time points, as the Ag derived from the UV-HSV inoculum presumably waned.

Furthermore, defective CD4⁺ T cell priming after UV-HSV inoculation was unlikely to be caused by differences in T cell recruitment to the LN, as HSV-specific CD4⁺ T cells appeared in the bLN shortly after inoculation with either form of the virus. When we cotransferred naive gDT-II together with control OT-II CD4⁺ T cells of irrelevant specificity, similar ratios of gDT-II over OT-II cells were observed 2 dpi in the bLN of mice inoculated with HSV-1 or UV-HSV (Fig. 3E, 3F). Additionally, similar frequencies of clustering gDT-II cells, defined as closely apposed (within 20 μm) like cells, were observed in the bLN 2 dpi, indicating comparable rates of gDT-II cell arrest and/or division on Ag-bearing DC regardless of the nature of the HSV inoculum (Fig. 3G, 3H). In summary, these data indicate that although UV-HSV inoculation delivered a reduced amount of Ag to the draining LN, these Ag levels were sufficient to attract and stimulate gDT-II cells during the first 3 dpi, suggesting that impaired effector differentiation was not solely attributable to a difference in Ag load.

The immune response to epicutaneous HSV-1 infection relies on DC-dependent trafficking of Ag to the draining LN, in part because of the confinement of infectious virus to the skin epithelium and later the skin-innervating neurons (37, 38). Following epicutaneous HSV-1 infection of mice, gDT-II cells are initially activated in the draining LN by migratory DC prior to interactions with resident CD8α⁺ DC, with this latter interaction being important for CD8⁺ T cell responses (20, 21, 39). We hypothesized that defective gDT-II cell priming following exposure to UV-HSV may have resulted from changes in the distribution of Ag across DC subsets within the bLN. Consequently, four major populations of DC found within cutaneous LN, namely LC, CD103⁺ dDC, and CD103⁻ dDC that have trafficked from the skin and LN-resident CD8a⁺ DC, were purified from bLN of mice that had been inoculated with HSV-1 or UV-HSV 2 d earlier and cultured with gDT-II cells (Fig. 4A). The purity of these isolated DC subsets was assessed prior to this experiment by extensive phenotyping in naive mice. As expected, CD8α⁺ DC displayed a “resident” phenotype as they were the only DC subset to display high CD11c and intermediate MHC II expression, differing from the classical CD11c^int MHC II^hi migratory DC phenotype displayed by LC, CD103⁻ dDC, and CD103⁺ dDC (Supplemental Fig. 1A, 1B) (40, 41). Furthermore, purified CD8α⁺ DC and CD103⁺ dDC populations were confirmed to be essentially homogenous, expressing high levels of XCR-1 with little to no expression of either CD11b or SIRPα, characteristic of cDC1 cells (Supplemental Fig. 1C) (3, 9, 42). By contrast, LC and CD103⁻ dDC mostly lacked XCR-1 and expressed intermediate and high levels of SIRPα, respectively (Supplemental Fig. 1C). Finally, both LC and CD103⁻ dDC also displayed heterogenous CD11b expression, particularly the CD103⁻ dDC subset, which contained equal proportions of CD11b⁻ and CD11b⁺ cells (Supplemental Fig. 1C). Following in vivo exposure to either the KOS or SC16 strain of live HSV-1 and consistent with previous findings, these CD103⁻ dDC as well as CD8α⁺ DC and CD103⁺ dDC were all able to elicit naive gDT-II cell proliferation in vitro, with the former two DC subsets observed to be the most potent stimulators of division (Fig. 4B, 4D) (20). CD8α⁺ DC and CD103⁺ dDC isolated from mice inoculated with UV-HSV, however, induced little proliferation of gDT-II cells, whereas CD103⁻ dDC were capable of stimulating gDT-II proliferation, albeit to a lesser degree than those isolated from infected animals (Fig. 4C, 4E). CD8α⁺ DC also appeared to be less activated than their counterparts from infected mice as they showed a more modest increase in CD40 expression after inoculation with UV-HSV, whereas the phenotypes of the migratory CD103⁻ dDC, CD103⁺ dDC, and LC were comparable between both groups (Supplemental Fig. 1D–F). Together, these data suggest that both LN-resident CD8α⁺ DC and CD103⁺ dDC had not acquired sufficient Ag in vivo following epicutaneous inoculation with UV-HSV to stimulate CD4⁺ T cell division.

Although differences in Ag load had the potential to impact the extent of T cell expansion and differentiation, they may have also impacted which DC populations were exposed to, and ultimately were capable of presenting, HSV-derived Ags. Consequently, we assessed the expansion and differentiation of CD4⁺ T cells in IRF8-deficient mice (Irf8^−/−), which lack CD8α⁺ DC and the related CD103⁺ dDC subset (Fig. 5A), allowing for an investigation of the role of these DC in CD4⁺ T cell priming in an environment replete with Ag (7, 43–45). HSV-1 infection of Irf8^−/− mice progressed in a similar manner to wild-type mice, with herpetic lesions appearing at 5 dpi (data not shown), although elevated viral titers were found in the infected skin of Irf8^−/−mice compared with control mice at this time (Supplemental Fig. 2A, 2B). Thus, in this model, HSV Ag is expected to be more abundant than after inoculation with nonreplicative UV-HSV but still cannot be presented by CD8α⁺ or CD103⁺ cDC1. Following infection of Irf8^−/−mice, adoptively transferred gDT-II cells divided to a similar extent to that observed in control mice, as evidenced by comparable CTV profiles and numbers of divided gDT-II cells (Fig. 5B, Supplemental Fig. 2C). Although T-bet expression was induced in gDT-II cells isolated from infected Irf8^−/− mice, which was not observed in wild-type mice inoculated with UV-HSV, the amount of T-bet was significantly reduced compared with that observed in gDT-II cells from infected wild-type controls (Fig. 5B, 5C). Although less pronounced, the proportion of CD25⁺ gDT-II cells was also lower in the Irf8^−/− mice (Fig. 5B, 5D). Critically, those gDT-II cells that expressed high levels of CD25 in infected Irf8^−/− mice possessed lower levels of T-bet than their counterparts from wild-type mice (Fig. 5E). Interestingly, this defect was not observed in mice deficient in IL-12, a Th1-polarizing cytokine produced by CD8α⁺ DC (5, 46–49). Indeed, comparable levels of T-bet and CD25 were expressed by gDT-II cells responding to epicutaneous HSV-1 infection in Il12p40^−/− and wild-type mice (Supplemental Fig. 3A–C).

To determine whether this partial defect in T-bet expression in gDT-II cells from IRF8-deficient mice resulted in impaired functional responses, the capacity of gDT-II cells isolated from infected wild-type and Irf8^−/− mice to produce IFN-γ and IL-2 following ex vivo restimulation with cognate peptide was assessed. Consistent with impaired Th1 differentiation, a significantly reduced proportion of gDT-II cells isolated from Irf8^−/− mice produced IFN-γ compared with those from control mice, whereas the impact on IL-2 production was more modest (Fig. 5F–H). Overall, these data indicate that following epicutaneous infection with HSV-1, CD4⁺ T cells stimulated in the absence of CD8α⁺ and CD103⁺ cDC1, and not simply in the absence of IL-12, exhibit significantly reduced levels of T-bet expression and, ultimately, impaired Th1 differentiation.

Finally, we reasoned that if Ag presentation by cDC1 was a requisite step for Th1 differentiation, then in the absence of active infection, direct targeting of Ag to these cells via the endocytic Clec9A receptor could potentially augment the generation of Th1 cells (50–53). Consequently, we employed a strategy whereby epicutaneous inoculation with UV-HSV was supplemented with Ag directly targeted to CD8α⁺ DC, as well as CD103⁺ dDC, using an anti-Clec9A Ab fusion construct containing glycoprotein D from HSV-1 (anti-Clec9A-gD) (29, 52, 54). This targeting approach, in the absence of polarizing adjuvants, has been shown to polarize CD4⁺ T cell differentiation toward T follicular helper and T regulatory cells (54, 55). Mice adoptively transferred with CTV-labeled gDT-II cells were first inoculated with UV-HSV via flank skin (Fig. 6A). Twenty-four hours later, when gDT-II cells were encountering Ag-bearing migrant dDC in the bLN, 2 μg of anti-Clec9A-gD or an isotype control Ig-gD construct (isotype-gD) was administered i.v. (21). Three days after receiving the initial UV-HSV inoculum, the extent of cell division and the expression of CD25 and T-bet by gDT-II cells within the bLN were assessed. As before, epicutaneous infection with HSV-1 induced robust proliferation and was accompanied by the induction of the expression of both CD25 and T-bet, whereas inoculation with UV-HSV alone failed to upregulate these molecules in gDT-II cells (Fig. 6B–E, Supplemental Fig. 4A–C). Although gDT-II cells isolated from mice that received UV-HSV and isotype-gD 24 h later had undergone a more robust proliferative response than mice that had received only UV-HSV, many cells did not upregulate CD25 and expressed low levels of T-bet (Fig. 6B–E, Supplemental Fig. 4A–C). A similar proliferative response was observed after administration of anti-Clec9A-gD to mice that had been inoculated with UV-HSV, with a higher proportion of gDT-II cells having undergone five or more divisions (Fig. 6B, 6C). Importantly, this was accompanied by dramatically increased expression of both CD25 and T-bet in gDT-II cells, significantly above that induced by inoculation with UV-HSV alone (Fig. 6B–E, Supplemental Fig. 4A–C). Supplementation with anti-Clec9A-gD elicited a higher proportion CD25⁺ gDT-II cells than in the UV-HSV + isotype-gD or anti-Clec9A-gD alone cohorts as well as significantly higher T-bet expression, particularly in the later divisions (Fig. 6D, 6E). Similar, albeit not significant, differences between these cohorts were observed when the numbers of CD25⁺ and T-bet⁺ gDT-II cells that had undergone at least one division were compared (Supplemental Fig. 4A–C). This phenotype most closely resembled that of gDT-II cells from infected wild-type mice and appeared to be the cumulative result of Ag presentation events within the bLN, as gDT-II cells isolated from the bLN of mice that received only anti-Clec9A-gD had not undergone as many rounds of division and had reduced levels of CD25 and T-bet expression relative to those observed in mice from the UV-HSV + anti-Clec9A-gD treatment cohort. Taken together, the data suggests that optimal Th1 differentiation requires the interactions between CD4⁺ T cells and cDC1, including LN-resident CD8α⁺ DC, that occur secondary to initial stimulation by migratory dDC.

We have shown that following administration of inactivated HSV, both CD4⁺ T cell expansion and functional differentiation are essentially defective. Most notably, inoculation with UV-HSV resulted in reduced levels of CD25 expression and impaired Th1 differentiation of gDT-II cells, phenotypic features that were evident as early as 3 dpi, suggesting that the priming environment was substantially different. Indeed, it appeared that Ag presentation differed not only in terms of the concentration of Ag within the bLN but also in the types of DC that contributed to the priming of CD4⁺ T cell responses. Contributions by CD103^– dDC and CD103⁺ dDC from the skin and CD8α⁺ DC resident within the bLN were essential for optimal Th1 responses to this pathogen. In line with this, supplementing UV-HSV skin inoculation with direct Ag delivery to cDC1 largely restored Th1 polarization.

CD4⁺ T cells have been reported to require multiple successive interactions with Ag-bearing DC to differentiate into effector cells (15–19). Indeed, Celli and colleagues (15) demonstrated that without a “second wave” of Ag presentation by DC allowing for continuing T cell/DC interactions during the inductive phase, upregulation of CD25, and acquisition of IFN-γ production were significantly diminished in CD4⁺ T cells. It was possible, therefore, that the impaired Th1 phenotype displayed by proliferating gDT-II cells in the bLN after priming with UV-HSV was due to a curtailed Ag presentation period. However, following inoculation of the skin with UV-HSV, there was still sufficient Ag to stimulate modest proliferation of naive gDT-II cells even 3–5 d later, despite the undoubtably lower abundance and briefer persistence of Ag in the priming LN. This suggests that at the very least, the impaired magnitude of the HSV-specific CD4⁺ T cell response and more particularly the impaired Th1 differentiation evident by day 3 is not purely the result of the lack of Ag within the bLN.

In addition to differences in Ag load, there are likely marked differences in the extent and nature of the innate immune response stimulated by the administration of UV-HSV compared with bona fide infection. Acute HSV-1 infection not only results in the generation of additional pathogen-associated molecular patterns that potentially signal through a range of pattern recognition receptors but also in the release of danger-associated molecular patterns from the lytic infection of the epithelium (56–60). Thus, the extent to which both Ag and danger signals are distributed within the draining LN may differ significantly after inoculation with the different forms of virus. Consistent with this, CD8α⁺ DC isolated from the bLN of mice given UV-HSV displayed a slightly less activated phenotype compared with their counterparts from the infection cohort. Moreover, in contrast to CD103⁻ dDC isolated from mice inoculated with UV-HSV, CD8α⁺ DC as well as CD103⁺ dDC from the same mice were unable to stimulate the proliferation of naive gDT-II cells in vitro. This suggests that they had not received sufficient Ag and/or had not been sufficiently exposed to maturation signals in vivo and that it was their lack of contribution to Ag presentation that caused defective Th1 differentiation. Consistent with this, following HSV-1 infection of mice that lacked cDC1 (Irf8^−/−), there were significant impairments in both the induction of T-bet and the generation of gDT-II cells capable of producing IFN-γ. Interestingly, the expression of CD25 on gDT-II cells from infected Irf8^−/− mice was more similar to that observed in wild-type mice, potentially owing to the levels of inflammation and Ag being more comparable.

We recently showed that following epicutaneous HSV-1 infection, CD4⁺ T cells interact with migrated DC from the skin prior to clustering with LN-resident XCR1⁺ CD8α⁺ DC during priming. However, the relative contributions of these DC subsets to the CD4⁺ T cell response, and in particular to T cell polarization, are not fully understood (21). The data presented in the current study indicates a functional consequence for sequential interactions of CD4⁺ T cells with different DC subtypes. When Ag presentation by CD8α⁺ DC and CD103⁺ dDC was compromised either as a result of the administration of UV-inactivated virus or a genetic ablation of cDC1, Th1 differentiation was impaired. Thus, although CD103^– dDC may provide an initial activation stimulus, this alone does not provide the full set of stimuli necessary for efficient Th1 differentiation. Indeed, Th1 differentiation in mice inoculated with UV-HSV was rescued when priming by CD103⁻ dDC was supplemented with Ag targeted to cDC1, implicating a requirement for cooperative priming by both migratory and LN-resident DC. Therefore, it can be surmised that cDC1 likely provide additional Th1 differentiation factors. Although the nature of these cues still requires elucidating, they appear to consist of more than simply the provision of IL-12, as Th1 differentiation remained intact in mice deficient in this cytokine. It is possible that a segregation of division and differentiation signals, between migrating DC from the skin and LN-resident DC, respectively, may act to restrict Th1 responses to only severe skin infections. Indeed, in the absence of sufficient Ag and inflammation to reach and activate LN-resident CD8α⁺ DC, relatively innocuous skin insults may result in curtailed and defective CD4⁺ T cell responses analogous to those seen after UV-HSV inoculation in this model. Such impairment would then presumably render these cells incapable of helping needless humoral and CD8⁺ T cell responses to these stimuli.

In summary, the data suggest that the generation of optimal Th1 CD4⁺ T cells in response to localized infections in which the pathogen is confined to the periphery relies on cooperation between CD103^– dDC and cDC1. Indeed, although the major contribution of the former subset is likely to convey Ag to the LN, resulting in early Ag presentation, the latter subsets are required for establishing and/or maintaining high levels of T-bet expression in CD4⁺ T cells. These findings have implications for future T cell vaccine design and highlight the importance of sustained Ag presentation and the involvement of multiple types of DC in the generation of Th1 responses to localized infections.

The authors have no financial conflicts of interest.