Dendritic Cells Are Dispensable for T Cell Priming and Control of Acute Lymphocytic Choriomeningitis Virus Infection

Lymphocytic choriomeningitis virus (LCMV) is an enveloped ssRNA virus and belongs to the group of arenaviruses that includes a variety of highly pathogenic hemorrhaging fever-causing viruses (1). LCMV is a noncytopathic virus and its main natural reservoir is small rodents, but it also infects humans and can cause severe symptoms. Several different laboratory strains of LCMV have been established and extensively characterized for their induction of protective immunity or viral persistence. Low-dose (200 PFU) infection of C57BL/6 mice with LCMV-WE or LCMV-Armstrong results in strong activation and expansion of CD8 T cells, the majority of which recognizes the immunodominant epitopes gp_33–41, nucleoprotein (NP)_396–404, and gp_276–286 (2). The activated CD8 T cells produce perforin, granzyme B, IFN-γ, TNF-α, and FasL, and thereby reduce the viral load to undetectable levels within 1–2 wk postinfection (3). Further, LCMV induces a strong CD4 T cell response, which is required for generation of neutralizing Abs, and these Abs appear critical to prevent secondary rise of LCMV titers months after the initial control by CD8 T cells (4). Initiation of the CD4 and CD8 T cell responses to LCMV is thought to be dependent on dendritic cells (DCs). This assumption is mainly based on experiments demonstrating that DCs are efficiently infected by LCMV (5) and efficient T cell priming could be achieved by transfer of LCMV peptide–pulsed DCs (6), targeting of LCMV peptides for specific uptake by DCs (7), or selective expression of LCMV peptides in DCs (8). Mice with selective expression of MHC class I (MHC-I) on DCs showed normal expansion of LCMV-specific CD8 T cells in the spleen in response to low-dose infection with LCMV-Docile (9). In contrast, diphtheria toxin (DT)–induced ablation of DCs in CD11c-DTR mice resulted in an impaired CD8 T cell response to LCMV (10). However, these results were not conclusive because later studies showed that DT treatment also ablated marginal zone metallophilic macrophages in the spleen (11, 12). These macrophages were found to be critical for limiting the spread of LCMV to other organs and to prevent exhaustion of CD8 T cells (13). Therefore, bystander toxicity of DT injection in CD11c-DTR mice made it impossible to define the relative role of DCs as APCs for priming the T cell response to LCMV. This raises the question whether DCs are indeed required to prime CD4 and CD8 T cells in response to LCMV infection.

Therefore, we investigated in this study the role of DCs for generation of efficient T and B cell response to LCMV using mice that constitutively lack DCs (ΔDC mice). ΔDC mice were generated by crossing CD11c-Cre mice to mice that express the DT α subunit (DTA) under control of a loxP-flanked stop cassette from the ubiquitously expressed Rosa26 locus. We have previously shown that ΔDC mice lack >95% of classical DCs and also the majority of plasmacytoid DCs and Langerhans cells (14). ΔDC mice mount an impaired Th2 response to helminth infection and a poor CD8 T cell responses to the model vaccine modified vaccinia virus Ankara encoding chicken OVA (14). In contrast, we observed in this study a normal CD4 and CD8 T cell response in wild-type (WT) and ΔDC mice. Low-dose LCMV-WE infection was efficiently cleared in both strains of mice. ΔDC mice also formed normal germinal centers (GCs), and serum levels of LCMV-specific Abs were not reduced compared with WT mice. Interestingly, depletion of macrophages in ΔDC or WT mice blunted the T cell response after LCMV infection. This indicates that macrophages are required and DCs are dispensable for induction of protective T cell immunity against acute LCMV infection.

Constitutively DC-ablated ∆DC mice (14), LCMV-specific CD8 TCR-transgenic P14_CD90.1 mice (15), LCMV-specific CD4 TCR-transgenic Smarta_CD90.1 mice (16), and B6_CD45.1 congenic mice (B6.SJL-Ptprca Pepcb/BoyJ) were maintained in the Franz-Penzoldt-Zentrum in Erlangen under specific pathogen-free conditions. C57BL/6J mice were purchased from Charles River Laboratories (Sulzfeld, Germany). If not stated otherwise, mice were infected with 200 PFU of LCMV-WE i.v. and analyzed at indicated time points. All experiments were performed in accordance with German animal protection law and European Union guidelines 86/809 and were approved by the Federal Government of Lower Franconia.

Single-cell suspensions of spleens or lymph nodes were generated by mechanical disruption, and erythrocytes were lysed with ACK buffer (0.15 M of NH₄Cl, 1 mM of KHO₃, 0.1 mM of Na₂ EDTA). Cells were preincubated with anti-CD16/CD32 mAb (clone 2.4G2; BioXCell, West Lebanon, NH) and stained with respective Abs. The following Abs were used for surface staining: PerCP-Cy5.5– or PE-labeled anti-CD11c (clone N418), PerCP-Cy5.5–labeled anti-CD4 (clone RM4-5), FITC- or allophycocyanin-labeled anti-CD8 (clone 53-6.7), PeCy7-labeled anti-CD62L (clone MEL-14), PE-labeled anti-F4/80 (clone BM8), eFluor660-labeled anti–GL-7 (clone GL-7), FITC- or eFluor450-labeled anti-CD45R (clone RA3-6B2), eFluor450-labeled anti–Gr-1 (clone RB6-8C5), and PE-labeled anti-CD45.1 (clone A20) were purchased from eBioscience (San Diego, CA). Allophycocyanin-Cy7–labeled anti–I-A/I-E (clone M5), Alexa Fluor 488–labeled anti-CD11b (clone M1/70), PeCy7-labeled anti-CD38 (clone 90), and PeCy7-labeled anti–PD-1 (clone RMP1-30) were purchased from Biolegend (San Diego, CA). Vioblue- or allophycocyanin-labeled anti-CD44 (clone IM7.8.1) were purchased from Miltenyi Biotec (Bergisch Gladbach, Germany), and PE-labeled anti-CXCR5 (clone 2G8) was from BD Biosciences (Heidelberg, Germany). For dextramer stainings, cells were washed with PBS + 5% FCS, incubated with 5 μl of dextramer per sample (gp33: KAVYNFATC; NP36: FQPQNGQFI; Immudex, Copenhagen, Denmark) for 10 min, and then the Ab mixture for surface staining was added. Tetramer (2 μg/ml, gp66: DIYKGVYQFKSV; National Institutes of Health Tetramer Core Facility) staining was performed in RPMI 1640 + 10% FCS (PAN-Biotech, Aidenbach, Germany). Cells were incubated with tetramer for 2 h at 37°C, washed, and stained with respective Abs. FITC-labeled anti-mouse IFN-γ (clone XMG1.2; Biolegend) and PE-labeled anti-mouse TNF-α (clone MP6-XT22; eBioscience) were used for intracellular staining after cells had been fixed with 4% paraformaldehyde and permeabilized with the Intracellular Staining Perm Wash Buffer (Biolegend) according to the manufacturer’s protocol. Dead cells were excluded by staining with DAPI (Sigma-Aldrich, St. Louis, MO), fixable viability dye allophycocyanin-eFluor780 (eBioscience), or fixable viability dye allophycocyanin-eFluor506 (eBioscience). Samples were acquired with FACSCanto II (BD Biosciences), and data were analyzed using FlowJo software (Tree Star, Ashland, OR).

Single-cell suspensions were restimulated with either 1 μg/ml gp33 (KAVYNFATM), gp61 (GLKGPDIYKGVYQFKSVEFD), or NP396 (FQPQNGQPI) peptide (JPT, Berlin, Germany) for 4 h. After 2 h, 10 μg/ml brefeldin A (Sigma-Aldrich) was added. IFN-γ and TNF-α production were measured by intracellular staining.

RNA was prepared with the RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s protocol. To transcribe the viral RNA genome, we reverse-transcribed 1 μg of total RNA into cDNA with an LCMV-specific reverse primer. Quantitative PCR was performed with SYBR Green I (Qiagen) and the following primer sequences: LCMV forward primer 5′-CATTCACCTGGACTTTGACAGACTC-3′ and LCMV reverse primer 5′-GCAACTGCTGTGTTCCCGAAAC-3′ under the following conditions: 95°C 30 s, 60°C 20 s, 72°C 20 s; 40 cycles. Copy numbers of LCMV genomes per gram spleen were determined using a plasmid containing 115 bp of LCMV GP gene.

A total of 5 × 10⁵ Smarta or 3 × 10⁵ P14 cells were either left untreated or stained with CellTrace Violet (5 μM; Molecular Probes, Eugene, OR) according to the manufacturer’s protocol and transferred to recipient mice 1 d before LCMV infection. Mice were analyzed on indicated days postinfection.

Splenocytes from naive B6 mice were stained with 5 μM of CellTrace Violet (Molecular Probes), loaded with 1 μM of gp33 peptide (JPT), and mixed with splenocytes from CD45.1 congenic mice loaded with 1 μM of m45 peptide of mouse CMV (JPT). The suspension of peptide-loaded cells was transferred into LCMV-infected mice at either day 8 or 28 postinfection. Analysis took place 90 min (day 8) or 12 h (day 28) after cell transfer. Percent specific killing was calculated as previously described (17).

Plates were coated with 1 μg/ml nucleoprotein of LCMV (Alpha Diagnostics, San Antonio, TX), blocked with PBS + 1% BSA, and incubated with sera overnight. Alkaline phosphatase–coupled anti-mouse IgG1 or anti-mouse IgG2c Abs and para-nitrophenylphosphate substrate (both Southern Biotech, Birmingham, AL) were used for detection. For detection of IFNs and IL12p40, commercial ELISA kits were used according to the manufacturer’s protocols (IFN-α and IFN-β: PBL Assay Science, Piscataway, NJ; IL12p40: R&D Systems, Minneapolis, MN). ELISA was measured at 405 nm with Multiskan FC multiplate photometer (Thermo Scientific).

To analyze the GCs, we fixed cryosections in acetone and rehydrated in PBS. GCs were stained with PE-labeled anti-mouse IgD (clone 11-26c; eBioscience), eFluor660-labeled anti-mouse CD45R (clone RA3-6B2; eBioscience), and with 25 μg/ml biotinylated peanut agglutinin (Vector Laboratories, Burlingame, CA) followed by incubation with DyLight488-labeled streptavidin (Biolegend). Nuclei were counterstained with DAPI.

Mann–Whitney U test and Student t test were performed with SigmaPlot (Systat Software, San Jose, CA) software, and p values <0.05 were considered statistically significant.

Low-dose LCMV-WE infection causes massive expansion of virus-specific CD8 T cells, which are required to reduce the viral load within 1 wk postinfection. To address the role of DCs for induction of this CD8 T cell response, we compared WT C57BL/6 mice with mice that constitutively lack DCs (ΔDC mice) because of CD11c-Cre–specific DTA expression (14). DC ablation appears to be very efficient (97% deletion) in ΔDC mice, and LCMV infection did not overcome this DC deficiency (Fig. 1A). The constitutive deletion of DCs in ΔDC mice did not result in a compensatory increase of other MHC-II–expressing cells types (Supplemental Fig. 1). Although CD11c is known to be expressed also on activated T cells (18), we did not observe a reduction of CD11c⁺ T cells in LCMV-infected ΔDC mice, which is surprising but might be explained by the lower expression level of CD11c in T cells as compared with DCs (Fig. 1B).

Type I IFNs (IFN-I) produced by conventional DCs (cDCs) rather than plasmacytoid DCs (pDCs) were reported to be critical for T cell priming after LCMV-Armstrong (3–6 × 10⁵ PFU) infection based on experiments where DCs had been depleted by injection of DT (19, 20). Therefore, it was important to investigate whether ΔDC mice show an impaired IFN-I response to low-dose (200 PFU) LCMV-WE infection. The IFN-I response was dominated by IFN-α, and no difference was observed in serum levels for this cytokine between WT and ΔDC mice. In contrast, serum levels of IFN-β and IL-12p40 were significantly reduced in ΔDC as compared with WT mice (Fig. 2).

Next, we analyzed the frequency and total number of LCMV-specific CD8 T cells in the spleen on day 8 after LCMV infection using fluorescently labeled H2-D^b dextramers loaded with the immunodominant LCMV peptides LCMV-gp_33–41 (gp33) and LCMV-NP_396–404 (NP396). We further restimulated splenocytes from LCMV-infected mice with gp33 and NP396 peptides to measure production of intracellular IFN-γ and TNF-α. Interestingly, the results of these experiments revealed no major differences between WT and ΔDC mice (Fig. 3A–D). We further determined the role of DCs for T cell priming at day 3 and 5 postinfection. Therefore, we performed adoptive transfers of CellTrace-labeled T cells from P14_CD90.1 mice in which most CD8 T cells express a LCMV-gp33-specific TCR. The proliferative response of transferred T cells was comparable between WT and ΔDC recipient mice (Fig. 3E).

To further determine the in vivo killing activity of CD8 T cells in LCMV-infected WT and ΔDC mice, we cotransferred gp33 peptide-pulsed target cells and target cells pulsed with an irrelevant peptide from mouse CMV (m45) on day 8 or 28 after LCMV infection. The specific killing efficiency was >80% and comparable between both strains of mice during the acute phase of infection (day 8) and during the memory phase (day 28) (Fig. 4A, 4B). We further measured the copy numbers of viral genomes in the spleen at different points in time after LCMV infection to determine the efficiency of viral clearance, which is largely dependent on CD8 T cells. In this study, we observed in both mouse strains the same viral loads early (day 4) after LCMV infection and a similar decline down to undetectable levels by day 28 postinfection (Fig. 4C).

These findings demonstrate that priming, cytokine production, killing capacity, and viral clearance by CD8 T cells are not impaired in ΔDC mice.

With the next set of experiments, we investigated the role of DCs for mounting an efficient CD4 T cell response against LCMV because CD4 T cells play an important role as helper cells for B cells to produce LCMV-specific Abs, which are required to control viral replication months postinfection (4). On day 8 postinfection, we restimulated splenocytes with LCMV-gp_61–80 peptide (gp61) and used fluorescently labeled I-A^b tetramers containing a shorter version of this peptide (gp66-Tet) to detect LCMV-specific CD4 T cells in the spleen. Unexpectedly, the CD4 T cell response to LCMV was not impaired in ΔDC mice because they had generated even more LCMV-specific CD4 T cells in comparison with WT mice (Fig. 5A, 5B). We further analyzed early T cell priming by adoptive transfer of CellTrace-labeled T cells from Smarta_CD90.1 mice in which most CD4 T cells express a transgenic TCR specific for LCMV-gp61 (16). Proliferation and expansion of transferred CD4 T cells was identical in WT and ΔDC recipients on days 3 and 5 postinfection (Fig. 5C). We followed expansion, decline, and activation status of transferred cells over 4 wk in the blood and observed no major differences between both strains of recipient mice (Fig. 5D, 5E). At day 28, we analyzed CD4 T memory cells in the spleen and observed the same number, activation status (CD44^hiCD62L^lo), and IFN-γ production of transferred T cells in WT and ΔDC mice (Fig. 5F, 5G).

To determine the memory T cells response of polyclonal CD4 and CD8 T cells, we first infected mice with 200 PFU of LCMV-WE and 4 wk later with 3 × 10⁵ PFU of LCMV-WE to elicit a memory T cell response in vivo. We analyzed the mice 4 d after secondary infection and observed a similar increase in the frequency of total CD44^hiCD62L^lo effector cells among CD4 or CD8 T cells in ΔDC and WT memory mice when compared with mice that received only the secondary infection (Fig. 6). The expansion of Ag-specific memory T cells was also comparable as determined by gp33-dextramer and gp66-tetramer staining (Fig. 6). After restimulation with LCMV-gp61 and LCMV-gp33 peptides, we detected the same frequency of IFN-γ⁺TNF-α⁺ T cells (Fig. 6).

This set of data indicates that DCs are not required for priming and expansion of LCMV-specific effector T cells nor for persistence and reactivation of memory T cells.

In addition to the CD4 and CD8 T cell response, we investigated the humoral response after LCMV infection. The frequency of GC B cells (B220⁺CD38^loGL-7^hi) and follicular Th (T_fh) cells (CXCR5^hiPD-1^hi) in naive ΔDC mice was higher as compared with WT mice (Fig. 7A, 7B). However, the LCMV-induced expansion of GC B cells and T_fh cells was comparable in both strains of mice (Fig. 7A, 7B). Histological analysis revealed efficient GC formation in both LCMV-infected WT and ΔDC mice (Fig. 7C). Consistent with the unimpaired GC response in LCMV-infected ΔDC mice, the LCMV-specific IgG1 and IgG2c serum levels were not reduced in ΔDC mice as compared with WT mice (Fig. 7D).

In this study, we investigated the role of DCs for priming of naive T cells and generation of a memory response during acute LCMV infection. Although it is well established that DCs can efficiently induce T cell immunity to LCMV, it remains unclear whether DCs are indeed required for T cell priming, acquisition of effector functions, and establishment of memory T cell populations. It was previously shown that selective and constitutive ablation of pDCs had no impact on CD4 and CD8 responses to acute LCMV infection but led to an impaired CD8 response to high-dose infection with the persistent LCMV strain Docile arguing for a critical role of pDCs as IFN producers to control persistent infections (21). Others have shown that cDCs are the main producers of IFN-I in response to acute LCMV infection, and cDC-derived IFN was required to mediate a CTL response to LCMV (19).

It has further been shown that DT-induced ablation of DCs in CD11cDTR mice results in a severely impaired CD8 T cell response to LCMV (10). However, using constitutively DC-depleted ΔDC mice, we found that DCs were not required for a normal cellular and humoral immune response upon acute low-dose LCMV infection. The IFN-α response, priming and expansion of CD8 and CD4 T cells, generation of memory T cells, as well as GC formation and Ab production, were comparable between ΔDC and WT mice.

How can the discrepancy between our results and the former studies be explained? Exogenous application of DT in CD11c-DTR mice efficiently depletes DCs but also causes depletion of the marginal zone metallophilic macrophages (11, 12), which are important to prevent spread of LCMV and T cell exhaustion (13). In contrast, the intrinsic DTA expression in DCs of ΔDC mice causes selective deletion of classical DCs, plasmacytoid DCs, and Langerhans cells, but not macrophages (12, 14).Therefore, ΔDC mice can be regarded as a more specific model to study the role of DCs for the immune response to LCMV.

One may argue that the few remaining DCs in ΔDC mice (∼3% of normal amounts) might be sufficient for T cell priming. Although we cannot completely rule out this possibility, we consider it highly unlikely because the chance for cognate interaction between DCs and T cells is severely reduced, and yet we observed no difference in T cell proliferation early (day 3 or 5) postinfection. Furthermore, we previously observed impaired T cell priming after modified vaccinia virus Ankara encoding chicken OVA immunization or helminth infection of ΔDC mice, demonstrating the important role of DCs in other immune responses (14). These results and our staining for MHC-II–expressing non-DC types also indicate that unimpaired T cell responses to LCMV in ΔDC mice are unlikely to be caused by compensatory expansion of other APCs. Further indication for a minor contribution of DCs for CD8 T cell priming during LCMV infection was provided by a study that showed that CD27/CD70 costimulation was required for CD8 T cell priming in transgenic mice that express LCMV epitopes only in DCs, whereas this costimulation pathway was not required for T cell priming upon LCMV infection of normal mice where Ag presentation is not restricted to DCs (5). Adoptively transferred P14 cells expand less efficiently after acute LCMV infection of mice in which only DCs can present MHC-I–restricted LCMV peptides as compared with WT recipient mice (9). This supports our findings that suggest that macrophages are more important for T cell priming than DCs in this infection model.

In support of our observations, CD169⁺ macrophages were shown to prime CTLs in ΔDC mice after adenoviral delivery of LCMV epitopes (12). An earlier study demonstrated impaired control of acute LCMV infection when macrophages had been depleted by administration of carrageenan (22). Others observed an impaired CTL response to LCMV and defective viral clearance after clodronate-induced ablation of marginal zone macrophages and metallophilic macrophages in the spleen (13). However, in this study, unimpaired in vitro CTL activity was found early (days 4 and 6), but not late (day 17), postinfection, suggesting that CD8 T cells get primed, but then acquire a state of exhaustion caused by uncontrolled viral spread.

The requirement of DCs as APCs might be linked to the replicative capacity of different viruses. It is known that immune responses to viruses with low replicative capacity in mice like influenza virus require costimulation (23) and are probably therefore more dependent on DCs as compared with the immune response to LCMV-WE, which replicates very fast. Furthermore, prolonged Ag stimulation circumvents the need for costimulation (24), leading to the conclusion that DCs play a minor role for priming T cell responses to viruses with high replicative capacity or long Ag persistence in other cell types.

Although we demonstrate in this study that CD4 and CD8 T cell responses to LCMV are dependent on macrophages, many other cell types get infected by LCMV and could serve as APCs. It was shown that B cell–deficient mice mount a poor primary CD4 response to LCMV (25) and loss of CD4 memory (26), which could point to a priming function of B cells or a structural function in the spleen. The latter study shows that membrane Ig transgenic mice, which have B cells but no LCMV-specific Abs, make a normal response. In a mouse model with B cell–restricted expression of MHC-II, Smarta cells expand after transfer and acute LCMV infection, and differentiation to T_fh occurs albeit at a lower level compared with WT mice (27), adding another hint to a priming function of B cells. Although in this study MHC-II–expressing B cells and DCs cooperate in CD4 T cell priming, this finding is still compatible with our results because we hypothesize that the DC-mediated effect could be taken over by macrophages in our system.

Furthermore, fibroblasts can also prime LCMV-specific CD8 T cells in lymphoid organs (28). The priming capability of the fibroblasts does not depend on costimulatory molecules but needs their migration to cytokine-rich lymphoid organs. In addition, it has been shown that bone marrow–derived professional APCs are not required for a CTL response after LCMV infection in parent (B6) → F1 (B6 × BALB/c) bone marrow chimeras (29).

Regarding the role of DCs for memory T cell formation, it is yet unclear whether DC-mediated priming is important to establish functional T cell memory to LCMV. It was recently shown that low- (1 μM) and high-dose (100 μM) LCMV-gp_33–41 peptide–pulsed DCs induce similar proliferation and immediate effector functions of CD8 T cells in vivo (30). But only DCs pulsed with high Ag load made stable T cell contacts and promoted transition of effector CD8 T cells to sustained memory T cells (30). However, by analyzing LCMV-infected ΔDC mice, we demonstrate in this study that DCs were not required to generate a population of CD8 memory T cells with immediate in vivo cytotoxic activity to transferred target cells. In addition, we observed normal expansion of both CD4 and CD8 memory T cells upon secondary infection at 4 wk after primary infection.

Taken together, we demonstrate that priming and acquisition of effector functions of CD4 and CD8 T cells upon acute low-dose LCMV infection occurs largely independent of DCs.

We thank B. Reizis for providing CD11c-Cre mice, the National Institutes of Health Tetramer Core Facility for providing LCMV-gp66_A-I^b Tetramers, K. Castiglione and L. Handl for technical assistance, M. Kirsch and L. Gundel for animal husbandry, and members of the Voehringer laboratory for critical comments.

The authors have no financial conflicts of interest.