Murine Intestinal Neonatal and Adult Dendritic Cell Subset Migration and Activation Characteristics in Response to Toll‐Like Receptor Stimulation Through MyD88

ABSTRACT

The initiation of adaptive immunity requires antigen presentation in immune inductive sites. Classical dendritic cells (cDC) sense peripheral immune response inducing cues and migrate to the tissue‐draining lymph nodes upon engagement, where they present acquired antigen to cognate T cells. Vaccines include adjuvants, which enable cDC activation and migration, both essential steps in the cascade leading to adaptive immunity. We here show that the expression of MyD88, the adapter molecule downstream of most toll‐like receptors (TLR), is essential for the migration of cDCs in response to several stimuli and that its restricted expression to the TLR‐sensing cDC subset is sufficient for the migration of that subset. As efficacious oral vaccines are primarily needed for use in newborns, we further quantify and characterise neonatal intestinal cDCs in five‐day‐old mice and show that they follow the same rules and patterns as we identified for cDCs in the adult intestines. Together, our data suggest strong conservation of immune‐inducing cues across ages and provide fundamental knowledge aiding adjuvant choice for neonatal vaccination strategies.

Despite their importance, dendritic cell subsets are not well characterised in neonates. We here provide fundamental information on the abundance and phenotype of small‐intestinal neonatal DCs and assess the role of MyD88 in neonatal and adult DC migration in response to several toll‐like receptor ligands.

1. Introduction

The event of birth marks an abrupt change from sterile conditions to life in an environment full of immunological triggers. The setup of lifelong immune homeostasis, including the establishment of a stable and beneficial microbiome, is the result of an intricate balance between protection from infection and learning to tolerate innocuous antigens. Enhanced occurrence of infections early in life as well as the finding that long‐term immune homeostasis depends on faithful priming during a critical time window before weaning suggest that there are substantial differences in the immune system between early life and adulthood.

Dendritic cells (DCs) are potent professional antigen‐presenting cells with crucial roles in the initiation of adaptive immunity due to their unique ability to migrate to draining lymph nodes (LNs) and to prime cognate naive T cells to peripherally acquired antigen [1, 2, 3, 4]. The two major subsets of classical DCs (cDC) are characterised by their transcription factor (TF) requirements and surface marker expression [5]. cDC type 1 (cDC1) depend on the TFs IRF8 and BATF3 and excel at cross‐presentation for the priming of CD8 T cell responses and at supporting Th1 responses towards intracellular pathogens and tumors [6, 7]. They have also been shown to be critical for presenting epithelial‐derived antigen in the mesenteric lymph nodes (mLNs) at steady state [8, 9] and for promoting oral tolerance [10]. Classical DC type 2 (cDC2) are more heterogeneous, but are generally implied in driving Th2 and Th17 responses to extracellular pathogens [11, 12, 13, 14], as well as IgA class switching in the Peyer's Patches towards both pathogenic and commensal bacteria [15, 16]. Recent work led to the sub‐division of cDC2 into Tcf4‐expressing pre‐DC2‐derived Notch2‐ and Bcl6‐dependent cDC2A and Klf4‐expressing pre‐cDC2‐derived cDC2B, whereby cDC2A are associated with the induction of Th17 and Tfh responses, while cDC2B drive Th2 immunity [17, 18, 19, 20, 21]. Whether cDC are absolutely required for the induction of Th17 has however been challenged recently [22].

Neonatal DCs are best characterised in the spleen, where they are relatively scarce and express lower levels of MHC class II and co‐stimulatory molecules at steady state, but where specific subsets resemble their adult counterparts in their expression of subset‐defining surface molecules [23]. Intestinal DCs are present at birth [24] and massively increase in response to infection with Cryptosporidium parvum , where they are critical for clearance [25]. In response to TLR9 engagement [26], antigenic targeting to CLEC9A or CLEC4A4 [27, 28] or combinatorial stimulation with several agonists [29], neonatal DCs can prime strong Th1 responses and even exceed adult cDC2 in their efficiency in priming Th17 responses under specific conditions [28].

Migration from the periphery to immune inductive sites is a key hallmark of cDC and essential for their ability to prime naïve T cells. In the adult intestines, DC migration at steady state and in response to the TLR7 ligand R848 or the TLR5 ligand flagellin critically depends on the DC‐intrinsic expression of the adaptor molecule MyD88 [30]. We and others have shown that sensing of type I IFN is required for optimal migration and activation by the responding cDC [31, 32]. TLRs differ in their expression pattern across ages [28, 33] and neonatal DCs show hypo‐responsiveness to type I IFN signalling [34]. Whether DC migration from the intestines to the draining mesenteric LNs differs between neonates and adults in response to TLR engagement has not been studied in detail before.

MyD88 is the key adapter protein required for downstream signalling events of the IL‐1 and IL‐18 receptors and all TLRs except TLR3 (and TLR4, which can signal through both MyD88 and TRIF) [35]. We here used conditional MyD88 knock‐out and knock‐in mice allowing for specific deletion or re‐expression of MyD88 in all CD11c‐expressing mononuclear phagocytes (MNPs) or only in cDC2 to in detail characterise the intrinsic requirements for DC accumulation and activation in the intestinal draining LNs in response to TLR stimulation across ages. We found that DC accumulation in the mLNs of adult mice depended more stringently on DC‐intrinsic MyD88 expression in response to R848‐mediated TLR7 stimulation than that in neonatal mice. MyD88 expression confined to cDC2 was sufficient for adult DC migration to flagellin, but only partly supported their migration in response to R848. Lastly, while cDC2 were required for Th17 priming in an adoptive transfer model using LPS‐mediated stimulation, mice in which MyD88 expression was confined to cDC2 were unable to support Th17 priming. Together, our findings contribute to a better characterisation of early life versus adult DC abundance, and migration kinetics and requirements. Additionally, they provide further insight into the role of cDC2 in the priming of Th17 immunity in adult mice.

2. Materials and Methods

2.1. Animals

All animals were maintained under specific pathogen–free conditions at Lund University Biomedical Center, the Technical University of Denmark (DTU) and the University of Calgary. C57Bl/6NRj mice were either purchased from Janvier or bred in‐house. The following strains were used: CD11c‐Cre.MyD88^{fl /fl} (B6.Cg‐Tg(Itgax‐cre)1‐1Reiz/J and B6.129P2(SJL)‐Myd88tm1Defr/J), huCD207‐DTA (B6.FVB‐Tg(CD207‐Dta)312Dhka/J), CD11c‐Cre.MyD88^{LSL /LSL} (B6.Cg‐Tg(Itgax‐cre)1‐1Reiz/J and ref. [36]), human‐Langerin‐Cre.MyD88^{LSL /LSL} [36, 37], XCR‐cre.YFP ( B6‐ Xcr1 ^tm2Ciphe and B6.129X1‐Gt(ROSA) 26Sor ^{tm1 (EYFP)Cos} /J), congenic CD45.1 OT‐II (B6.Cg‐Tg(TcraTcrb)425Cbn/J, B6.SJL‐Ptprc^a Pepc^b/BoyJ background) and BDCA2‐DTR (C57BL/6‐Tg(CLEC4C‐HBEGF)956Cln/J). For floxed models, cre was bred heterozygously and cre‐negative mice served as wt (for fl/fl models) or full KO (for LSL/LSL models) control. For the latter, age‐and sex‐matched C57Bl/6 non‐littermates were used as wt controls. All experiments were conducted with approval from the respective national animal welfare authorities.

2.2. Immunisation

Five‐day‐old neonatal mice or adult mice (12–16 weeks old) were orally immunised with 2 μg and 20 μg of Resiquimod (R848) in PBS, respectively, or intraperitoneally with 20 μg FliC [38] or 100 μg poly(I:C) (Sigma‐Aldrich). Unless otherwise specified, mice were analysed at 15 h post‐immunisation. BDCA2‐DTR mice received either PBS or 200 ng diphtheria toxin (DTA; EMD Millipore, Corynebacterium diphtheriae , unnicked) in PBS on day −1 and day 0 intraperitoneally.

2.3. Adoptive Transfer of T Cells

Naïve CD4⁺ T cells were isolated from the spleen and mesenteric lymph nodes (mLN) of CD45.1 OT‐II mice with the Dynabeads Untouched Mouse CD4 Cells kit or the Naïve CD4⁺ T Cell Isolation kit from STEMCELL Technologies following the manufacturer's protocol. A total of 1 × 10 [6] naïve CD4⁺ T cells were transferred into recipient mice via intravenous injection. 24 h post‐transfer, mice were intraperitoneally injected with 0.5 mg ovalbumin (OVA; Grade V, Sigma‐Aldrich), 25 μg anti‐CD40 and 20 μg LPS. To prevent lymphocyte egress from lymph nodes, 20 μg FTY20 was administered on days 1 and 3 following immunisation. Mice were sacrificed 4 days after immunisation, and mLNs were harvested for analysis.

2.4. Cell Isolation

Single‐cell suspensions were prepared from mLNs and spleens by mechanical disruption through 70 μm cell strainers. Cells were resuspended in FACS buffer (PBS supplemented with 2% FCS and 2 mM EDTA) and subsequently subjected to restimulation and/or staining for flow cytometry. Small‐intestinal single cells were obtained by flushing out the intestinal content with cold HBSS supplemented with 15 mM HEPES. The intestine was longitudinally opened and cut into small pieces. Epithelial cells were removed by incubating the tissue for 15 min at 37°C with 2 mM EDTA in HBSS supplemented with 10% FCS. This was followed by vigorous shaking and filtering using a nylon mesh. The EDTA washing was repeated twice for the younger mice (before weaning) and 3 times for adult mice, and the remaining tissue was digested in RPMI medium (Gibco cat#11875093) containing 10% FBS, 0.3 Wünsch/mL Liberase TM (Roche cat# 05401119001) and 30 μg/mL DNase I (Roche cat#10104159001) on a magnetic stirrer for 15–20 min at 37°C. The digested cell suspensions were filtered through a 100 μm strainer (Fisher Scientific) and lymphocytes were enriched by density gradient centrifugation with 40%/70% Percoll (Cytiva #17089101).

2.5. In Vitro T Cell Stimulation

A total of 1 × 10⁷ mesenteric lymph node (mLN) cells were incubated in complete medium supplemented with PMA (25 ng/mL; Sigma‐Aldrich) and ionomycin (500 ng/mL; Sigma‐Aldrich) for 4 h at 37°C in 5% CO₂. Golgi Stop (1 μL; BD Biosciences) was added after 1 h to inhibit protein transport.

2.6. Flow Cytometry

For dendritic cell (DC) staining, non‐specific binding was blocked using 10% rat serum and rat anti‐mouse CD16/CD32 Fc block for 20 min at 4°C. Dendritic cells were subsequently identified using the following antibodies. AF700 anti‐CD3 (17A2), AF700 anti‐CD19 (1D3), AF700 anti‐NK.1.1 (PK136), AF700 anti‐B220 (RA3‐6B2), APC anti‐CD64 (X54‐5/7.1), BV510 anti‐CD45 (30‐F11), PE‐Cy7 anti‐CD11c (N418), BV650 anti‐MHC‐II (M5/114.15.2), PE anti‐XCR1 (ZET), APC‐Cy7 anti‐CD11b (M1/70) and FITC anti‐CD86 (GL1). Cell aggregates were excluded based on their FSC‐A and FSC‐W, and dead cells were excluded based on positive staining for propidium iodide (Sigma‐Aldrich).

For steady‐state stains shown in Figure 1 only, single cell suspensions were stained with LIVE/DEAD Blue fixable viability dye (Invitrogen) together with CD16/CD32 FC block for 25 min on ice followed by staining with BUV395 anti‐CD45 (30‐F11), BUV737 anti‐CD11b (M1/70), FITC anti‐Sirpα (P84), BV421 anti‐F4/80 (BM8), BV785 anti‐CD103 (2e7), PE anti‐CD101 (Moushi101), PE‐Dazzle anti‐CD11c (N418), PE‐CY5 anti‐CD3 (145‐2C11), CD19 (eBio1D3), B220 (RA3‐6B2), NK1.1 (PK136), PE‐CY7 anti‐CD64 (X54‐5/7.1), APC anti‐XCR1 (ZET) and APC‐Fire 810 anti‐MHCII (M5/114.15.2).

FIGURE 1.

Dendritic cell occurrence and phenotype in neonatal versus adult mice at steady state. (A) Representative flow cytometry plots of DCs and subsets from the lamina propria of 5‐day‐old, 10‐day‐old, 20‐day‐old and adult (11–15 weeks) mice in steady‐state conditions. Each symbol represents an individual mouse. DCs were pre‐gated on single, live, lineage‐negative (CD3, CD19, B220, NK1.1), F4/80 and CD64 negative cells. Data is representative of 2 independent experiments with 2–3 mice each for ages of 5, 10 and 20 days and 4 experiments with 3 mice each for adult timepoints. (B) Representative histograms showing various cDC1 and cDC2 markers, pre‐gated on XCR1⁺ (top, shades of green) and CD11b⁺XCR1⁻ (shades of orange) (gates as indicated in Figure 1A) in the small‐intestinal lamina propria of mice of the indicated ages at steady‐state conditions. (C) Fluorescence microscopy image of small intestine lamina propria from 6‐day‐old neonatal mice carrying an eYFP reporter for XCR1, showing cDC1 (CD11c⁺MHC II⁺XCR1⁺CD64⁻) and cDC2 (CD11c⁺MHC II⁺CD101⁺CD64⁻) at steady state. EYFP staining was intensified using an αGFP antibody raised in chicken and an α‐chicken‐A488 secondary antibody. White arrows point at cDC1, red arrows at cDC2. (D) Absolute number of total DCs (CD45⁺lin⁻CD64⁻F4/80⁻CD11c⁺MHCII⁺), cDC1 (CD103⁺CD11b⁻), double‐positive cDC2 (CD103⁺CD11b⁺XCR1⁻) and CD11b single‐positive cDC2 (CD103⁻CD11b⁺XCR1⁻) subsets in the small‐intestinal lamina propria at indicated postnatal age. Data is representative of 2 independent experiments with 2–3 mice each for ages of 5, 10 and 20 days and 4 experiments with 3 mice each for adult timepoints. Each symbol represents an individual mouse. Bars indicate mean +/− SD. One‐way ANOVA using Brown–Forsythe and Welch (*p < 0.05, **p < 0.005, ***p < 0.0005, ****p < 0.0001).

For identification of Th17 CD4⁺ T cells, non‐specific binding was blocked with 10% rat serum and anti‐mouse CD16/CD32 Fc block for 20 min at 4°C. Cells were then stained with the following surface antibodies: PE anti‐CD8α (53–6.7), BV786 anti‐CD45.1(A20), A700 anti‐CD45.2 (104), PE‐CF594 anti‐TCRβ (H57‐597), APC‐Cy7 anti‐CD4 (RM4‐5), PE‐CY7 anti‐CD44 (IM7), and APC‐Cy7 anti‐CD62L (MEL‐14). The cells were then fixed and permeabilised using the Foxp3/Transcription Factor Staining Buffer Set (eBioscience) according to the manufacturer's instructions, followed by intracellular staining using FITC anti‐mouse IL‐17A (TC11‐18H10.1). Cell aggregates were excluded based on their FSC‐A and FSC‐W, and dead cells were excluded using LIVE/DEAD Fixable Aqua Dead Cell Stain Kit (Thermo Fisher Scientific).

2.7. Immunofluorescence Staining of Tissue Sections

Tissues were fixed for 1 h in Antigen Fix (3% PFA; Diapath), followed by overnight dehydration in 35% sucrose and embedding in Tissue‐Tek OCT. Embedded tissues were sectioned at 20 μm using a cryostat (Leica CM1950).

Cryostat sections were blocked overnight at 4°C in PBS containing 0.5% saponin, 2% FCS, 2% BSA and 2% donkey serum. For immunostaining, tissues were incubated overnight with the indicated primary antibodies, followed by 1‐h incubation with the appropriate secondary antibodies. When tertiary antibodies were used, an additional saturation step was included to prevent cross‐reactivity with secondary antibodies.

Following staining, tissue sections were washed and mounted in ProLong Gold (Molecular Probes) containing Sytox Blue (ThermoFisher) and imaged using a Zeiss LSM780 confocal microscope in online fingerprint mode. The following antibodies were used to identify the respective cell types in the tissue: Primary antibodies: Chicken anti‐GFP (polyclonal), mouse anti‐CD64 (X54‐5/7.1), rat anti‐CD101 (307707), hamster anti‐CD11c (N418), anti‐CD4 (RM4‐5) and anti‐MHCII (M5/114) (all from Invitrogen); secondary antibodies: Anti‐Chicken‐AF488 (ThermoFisher), Anti‐Rat‐Cy3 and Anti‐Hamster‐AF594 (Jackson ImmunoResearch). After this, the stained tissue sections were visualised with a Zeiss LSM 780 confocal microscope using online fingerprint mode. Images were analysed using Adobe Photoshop 2020.

2.8. Statistical Analysis

Statistical analysis was performed by Mann–Whitney U test or ANOVA with Tukey's post‐hoc test using GraphPad Prism software (GraphPad) throughout the manuscript except Figure 1D, for which Brown–Forsythe parametric one‐way ANOVA was used.

3. Results

3.1. Intestinal DCs Are Present in the Neonatal Intestines With Subset Differences in Kinetics and Phenotypes

Dendritic cells arise from the bone marrow, seed the periphery starting at birth and are constantly replenished throughout life. We first set out to phenotypically compare subsets of DCs in the small intestine and track their appearance in early life. At day 5 post birth, the earliest timepoint investigated in our study, we identified a clear population of CD11c⁺MHCII⁺ cells, which stained negative for the macrophage markers CD64 and F4/80 (Figure 1A, for pre‐gating see Figure S1). These cells were enriched for XCR1⁺ cells, a phenotype characteristic for cDC1. We detected very few XCR1⁻CD11b⁻ cells within the intestinal cDC population at neonatal day 5, suggesting that the XCR1⁻ phenotype of fetal/neonatal cDC1 recently described in the lung might be specific to that organ, or disappear from the intestines at an earlier age [39]. The neonatal small intestine also contained a population expressing CD11b⁺, but not XCR1, which did not express CD103 to the same extent as found in the adult small intestine. Instead, the CD103⁺CD11b⁺ population of cDC2 gradually increased over the timepoints sampled (d5, d10, d20 and 11–15 weeks post birth). Despite the low CD103 expression in early life small‐intestinal cDCs, many however expressed CD101, a marker for CD103⁺CD11b⁺ cDC2 in the adult small intestine (Figure 1A,B). To better understand whether neonatal DC subsets indeed represented cDC1 and cDC2, we directly compared cDC1 and cDC2 associated surface markers on XCR1⁺ versus CD11b⁺XCR1⁻ cDCs. We found that XCR1⁺ cDCs expressed similarly high levels of XCR1 and CD103 regardless of age and were negative for the cDC2 markers Sirpα and CD101, leading us to conclude that XCR1 is a reliable cDC1 marker also in the neonatal setting. Curiously, XCR1⁺ cells express higher levels of CD11b at the d5 and d10 timepoints, which however does not reach the level of expression detected in XCR1⁻ DCs (Figure 1B). The CD11b⁺XCR1⁻ cDC compartment expressed similar levels of CD11b throughout the timepoints sampled. Within that population, the contribution of cells co‐expressing Sirpα and CD101 gradually increased with age, but the expression level of these markers per cell did not change (Figure 1B). Lastly, to confirm the presence of cDC subsets in the neonatal small‐intestinal lamina propria, we stained small‐intestinal sections from 5‐day‐old mice carrying eYFP as a fluorescent reporter under the control of the XCR1 promotor (for cDC1) with CD11c (pan DCs), CD101 (as a cDC2 marker) and CD64 (to exclude macrophages) (Figure 1C). Together, we conclude that neonatal intestines at day 5 post birth contain both cDC1 (XCR1⁺ DCs) and cDC2 (CD11b⁺XCR1⁻ DCs), with the latter showing variable expression of cDC2 markers, potentially reflecting differences in maturation or subset heterogeneity across ages. Both cDC1 and cDC2 subsets were relatively low in numbers in the SILP 5 days after birth. cDC1 significantly increased in abundance by day 10 of age, reaching adult numbers already at this timepoint. Instead, cDC2 gradually increased with age, showing a significant increase between day 20 and adulthood (Figure 1D). This is in line with previous findings showing that cDC2, but not cDC1, are reduced in Peyer's Patches (PPs) during early life [40]. Of note, we did not remove PPs from the small‐intestinal cell preparations in any of our samples taken from mice pre‐weaning.

3.2. Neonatal Intestinal DCs Efficiently Migrate to the Mesenteric Lymph Nodes After R848 Gavage

Migration to the draining lymphoid tissues for antigen presentation is a prerequisite for peripheral DC function. Steady‐state migration is enhanced significantly upon DC activation leading to upregulation of CCR7 for the sensing of the CCL19/CCL21 gradient governing homing to lymphoid tissues [41]. To test the migration capacity of neonatal DCs, we orally gavaged 5 days old pups with R848, a synthetic ligand for TLR7 known to induce strong DC migration in adult mice [31]. By assessing the accumulation of DCs in the draining mesenteric LNs, we found that neonatal DCs readily migrated in response to R848 (Figure 2A). Neonatal cDC1 and cDC2 gradually accumulated in the mLN, where their contribution to overall cellularity peaked at 18–20 h. We also measured a gradual increase in the overall expression of the co‐stimulatory molecule CD86 on DCs in the mLNs as a sign of activation (Figure 2B). In contrast, CD86 expression peaked earlier and then decreased again in DCs within the SILP, presumably reflecting the egress of activated DCs (Figure 2C). Of note, activation of DCs was also prominent in the spleen upon oral R848 stimulation, suggesting that DC activation may not depend on cell‐intrinsic sensing of R848 and instead is mediated by soluble factors inducing a systemic effect to R848 feeding (Figure 2D). This is in line with our previous findings that DCs readily migrate in response to poly(I:C) stimulation if activated in‐trans [32], and with data from Yrlid et al. [31], describing an essential role for TNFα and type I IFN production by pDCs for the migration of cDCs in response to R848 feeding.

FIGURE 2.

Neonatal dendritic cells exhibit migratory capacity upon R848 stimulation. Five‐day‐old neonatal mice were orally administered 2 μg of R848 in PBS, and dendritic cell (DC) migration to the mesenteric lymph nodes (mLN) was analysed at the indicated times post gavage. (A) Frequency of total DCs (CD45⁺CD64⁻CD11c⁺MHC II⁺), cDC1 (CD103⁺CD11b⁻), double‐positive cDC2 (CD103⁺CD11b⁺) and CD11b single‐positive cDC2 (CD103⁻CD11b⁺) among total haematopoietic cells (CD45⁺) at the indicated time points. Graphs show pooled data from multiple experiments: 0 h (5 experiments, 1–3 mice per experiment), 2 h (1 experiment, 4 mice), 4 h (1 experiment, 3 mice), 9 h (1 experiment, 3 mice), 16 h (2 experiments, 3 mice each), 18 h (2 experiments, 3–4 mice each), 20 h (2 experiments, 4–6 mice each), 24 h (2 experiments, 3–4 mice each), 36 h (1 experiment, 4 mice) and 48 h (1 experiment, 4 mice). Each symbol represents an individual mouse. (B) Fold change in CD86 median fluorescence intensity (MFI) relative to unstimulated mice in the mLN at the indicated time points. (C) Fold change in CD86 MFI relative to unstimulated mice in the SILP at the indicated time points. (B, C) Graphs show pooled data from multiple experiments: 2 h (1 experiment, 4 mice), 4 h (1 experiment, 3 mice), 9 h (1 experiment, 3 mice), 16 h (1 experiment, 3 mice), 18 h (1 experiment, 4 mice), 20 h (1 experiment, 4 mice), 24 h (1 experiment, 4 mice), 36 h (1 experiment, 4 mice) and 48 h (1 experiment, 4 mice). Each symbol represents an individual mouse. (D) Fold change in CD86 MFI relative to unstimulated mice in the spleen at the indicated time points. Graphs show pooled data from multiple experiments: 4 h (1 experiment, 3 mice), 9 h (1 experiment, 3 mice), 16 h (2 experiments, 3 mice each), 18 h (1 experiment, 3 mice), 20 h (2 experiments, 3 mice each), 24 h (1 experiment, 3 mice), 36 h (1 experiment, 4 mice) and 48 h (1 experiment, 4 mice). Each symbol represents an individual mouse.

3.3. MyD88 In CD11c‐Expressing Cells Is Required and Sufficient for DC Migration in Response to R848 Across Ages

Most TLRs signal through MyD88, and this adapter has been shown to be essential for the migration of DCs in response to R848 in adults [30]. Indeed, mice in which all CD11c‐expressing cells lacked MyD88 showed no DC migration in response to R848 (Figure 3A). In contrast, neonatal cDC1 (and potentially cDC2, the latter much less abundant at this age, prohibiting firm conclusions) showed a partial increase in the mLN in response to oral R848 treatment in the neonates that lacked MyD88 in the CD11c‐expressing cellular compartment (Figure 3B). This suggests that at an early age, CD11c‐negative cells can sense R848 and support cDC accumulation in the mLN.

FIGURE 3.

MyD88 signalling in CD11c‐expressing cells is required for dendritic cell migration to the mesenteric lymph nodes upon R848 stimulation. (A) Adult mice (12–16 weeks) and (B) 5 days old neonatal mice of the indicated genotypes (CD11c‐Cre.MyD88^fl/fl or MyD88^fl/fl littermates (WT control)) were orally administered 20 μg and 2 μg of R848 in PBS, respectively. Mesenteric lymph node DCs (live, lineage‐negative, CD11c⁺MHCII⁺) were analysed 15 h after immunisation for abundance of cDC1 (CD103⁺CD11b⁻) and cDC2 (CD103⁺CD11b⁺). Data are pooled from three independent experiments, with 3–4 mice per group in each experiment. Each symbol represents an individual mouse; black = PBS control, orange = R848‐treated mice. (C) Adult mice (12–16 weeks) and (D) five‐day‐old neonatal mice of the indicated genotypes (CD11c‐Cre.MyD88^LSL/LSL [CD11c.MyD88^ON], MyD88^LSL/LSL [KO] littermates and C57Bl/6 unrelated WT controls) were orally administered 20 μg and 2 μg of R848 in PBS, respectively. Mesenteric lymph node DCs (live, lineage‐negative, CD11c⁺MHCII⁺) were analysed 15 h after immunisation for abundance of cDC1 (CD103⁺CD11b⁻) and cDC2 (CD103⁺CD11b⁺). Data are pooled from three independent experiments, with 2–6 mice per group for adults and 3–5 mice per group for neonates, except for the PBS‐treated neonatal controls, which were tested once with 3 mice per genotype (indicated on the graph). Each symbol represents an individual mouse; black = PBS control, orange = R848‐treated mice. (E) Adult mice (12–16 weeks) of the indicated genotypes (CD11c‐Cre.MyD88^LSL/LSL [CD11c.MyD88^ON], MyD88^LSL/LSL [KO] littermates and C57Bl/6 unrelated WT controls) were administered 100 μg poly(I:C) intraperitoneally (i.p.) in PBS. Mesenteric lymph node DCs (live, lineage‐negative, CD11c⁺MHCII⁺) were analysed 15 h after immunisation for abundance of cDC1 (CD103⁺CD11b⁻) and cDC2 (CD103⁺CD11b⁺). Data are pooled from two independent experiments, with 3–5 mice per group. Each symbol represents an individual mouse; black = PBS control, blue = poly(I:C)‐treated mice. Statistical analyses were performed using one‐way ANOVA with Tukey's post hoc, (*p < 0.05, **p < 0.005, ***p < 0.0005, ****p < 0.0001).

Since MyD88 is broadly expressed, it is theoretically possible that DC migration, in addition to expression in cDC11c‐positive cells, requires additional signals derived from the periphery. To assess whether MyD88 expression in the CD11c‐positive compartment alone was sufficient for DC migration, we used CD11c.MyD88^LSL/LSL mice in which MyD88 is re‐expressed within the CD11c‐positive compartment in the endogenous locus on an otherwise MyD88‐deficient background [36]. Migration of both cDC1 and cDC2 was completely restored in response to oral R848 in both adult and neonatal mice (Figure 3C,D). For neonatal experiments in this transgenic model, MyD88^KO mice were CD11c.cre‐negative MyD88^LSL/LSL littermates from CD11c.cre‐positive MyD88^LSL/LSL (CD11c.MyD88^ON) pups; wildtype control littermates ubiquitously expressing MyD88 were hence not available. cDC numbers in the mLNs from R848‐treated cre‐negative MyD88^fl/fl controls were however comparable to those in CD11c.MyD88^ON pups (Figure 3B). One MyD88^KO pup curiously showed wildtype‐level migration (Figure 3D), and we suspect this to be due to a genotyping failure as cre‐expression in the CD11c.cre x MyD88^LSL/LSL line leads to frequent occurrence of germline deletion of the LSL allele, reverting the system back to a wildtype scenario.

To rule out general defects of cDCs in the absence of MyD88, we next checked migration of DCs in response to poly(I:C) in the MyD88^ON model, which we previously showed to critically depend on TLR3/TRIF signalling within the DC compartment [32]. Particularly cDC1 migrated even better in CD11c.MyD88^LSL/LSL compared to wt mice (Figure 3E), presumably due to additional maturation signals provided by a weakened barrier in mice devoid of MyD88 signalling in non‐CD11c‐expressing cells.

Together, these data suggest that MyD88 expression within the CD11c‐positive cellular compartment is sufficient for DC migration and that the early immune activation events in response to TLR7 stimulation are generally conserved across neonates and adults.

3.4. MyD88 Expression in cDC2 Is Sufficient for Migration in Response to FliC, but Not in Response to R848

Studying poly(I:C)‐induced DC migration mediated via the TLR3/TRIF pathway, we have previously found that migration and activation can occur in trans, meaning that cell‐intrinsic expression of TLR3 was not required [32]. To study whether this was also the case for cDC2 in the context of MyD88‐mediated activation pathways, we bred mice in which only intestinal cDC2 expressed MyD88 on an otherwise MyD88‐deficient background. To achieve this, we used huCD207‐cre mice, which faithfully express cre in migratory small‐intestinal cDC2 in addition to Langerhans cells [42].

TLR7‐expression is more highly expressed in plasmacytoid DCs (pDCs), while cDC2 express high levels of TLR5, the receptor for flagellin [43, 44]. Interestingly, oral gavage of huCD207.MyD88^ON mice with R848 failed to induce substantial cDC2 migration (Figure 4A), indicating that cDC2 might not express sufficient TLR7 to migrate in response to R848 gavage or that additional signals are required for their migration in addition to in‐cis activation. R848 gavage in CD11c.MyD88^ON mice, in which all CD11c⁺ cells including pDCs re‐express MyD88, led to efficient cDC2 migration (Figure 3C,D) and Yrlid et al. previously published that pDCs facilitated DC migration upon R848 feeding [31]. To test whether pDCs indeed provided the signals required for cDC2 migration in response to R848, we used BDCA2‐DTR mice that allow for in vivo depletion of pDCs by diphtheria toxin (DT) injection. Some mouse models using the DTR transgene present with a general LN hypotrophy [45], and we found this to also be the case in BDCA2‐DTR mice (data not shown). We therefore used PBS‐treated transgenic controls for DT‐treated transgenic experimental groups throughout. Partial depletion of pDCs prior to R848 gavage (Figure 4B) led to significantly blunted migration of cDC1, but not cDC2, upon R848 gavage (Figure 4C). Together, these data show that MyD88 expression in intestinal cDC2 alone is insufficient to induce migration in the context of R848 gavage, but that MyD88 expression in total CD11c‐expressing cells recovers cDC2 migration in a manner insensitive to partial pDC depletion. This is in line with our previous findings that cDC2 are less dependent on type I IFN than cDC1 [32], suggesting that migration can be maintained under sub‐optimal conditions.

FIGURE 4.

MyD88 expression in cDC2 is sufficient to drive DC migration to the mLN in response to FliC but not R848. (A) Adult mice (12–16 weeks) of the indicated genotypes (human‐Langerin‐cre.MyD88^LSL/LSL [CD207.MyD88^ON], MyD88^LSL/LSL [KO] littermates and C57Bl/6 unrelated WT [controls]) were orally administered 20 μg R848 in PBS. Mesenteric lymph node DCs (live, lineage‐negative, CD11c⁺MHCII⁺) were analysed 15 h later for cDC1 (CD103⁺CD11b⁻) and cDC2 (CD103⁺CD11b⁺) abundance. Data are pooled from two independent experiments (2–6 mice/group). Each symbol represents an individual mouse; black = PBS control, orange = R848. (B) BDCA2‐DTR mice were orally administered 20 μg R848 in PBS after receiving either 200 ng diphtheria toxin or PBS on days −1 and 0 intraperitoneally. Mesenteric lymph node DCs (live, lineage‐negative, CD11c⁺MHCII⁺) were analysed 15 h later for cDC1 and cDC2 abundance. Data are pooled from two independent experiments (4 mice/group). Each symbol represents an individual mouse. (C) Adult mice (12–16 weeks) of the indicated genotypes (human‐Langerin‐cre.MyD88^LSL/LSL [CD207.MyD88^ON], MyD88^LSL/LSL [KO] littermates and C57Bl/6 unrelated WT [controls]) were administered 20 μg FliC in PBS intraperitoneally. Mesenteric lymph nodes were analysed 15 h later for cDC1 and cDC2 abundance. Data are pooled from two experiments (2–5 mice/group for FliC‐treated) and one experiment (3–4 mice/group for PBS controls). Each symbol represents an individual mouse; black = PBS control; green = FliC. Statistical analyses were performed using one‐way ANOVA with Tukey's post hoc, (*p < 0.05, **p < 0.005, ***p < 0.0005, ****p < 0.0001).

Lastly, to test whether lack of cDC2 migration in huCD207.MyD88^ON mice in response to R848 might be due to insufficient TLR7 signalling in cDC2, we interrogated their migration in response to FliC, engaging with TLR5, which is highly expressed on intestinal cDC2. In this scenario, cDC2 migrated normally, showing that TLR5 engagement on cDC2 alone is sufficient for their migration (Figure 4D). FliC did not induce significant migration of cDC1 even in wildtype conditions. Unfortunately, this experimental setup could not be faithfully tested in neonates, as the MyD88‐LSL locus shows high sensitivity to huCD207.cre‐mediated adverse germline deletion, prohibiting the generation of sufficient neonates for meaningful statistics in a scenario in which genotype could not be determined before immunisation. Nevertheless, we conclude that cDC2 readily migrate in adult and neonatal mice in response to R848 and FliC, the former requiring accessory CD11c⁺ cells capable of MyD88‐mediated TLR signalling for optimal migration, the latter only requiring MyD88 activity in the cDC2 compartment.

3.5. cDC2 Are Required, but Not Sufficient for Th17 Induction in a CD4 T Cell Transfer Model

cDC2 have previously been shown to potently induce intestinal Th17 immunity [11, 13], but recent studies challenge their relative importance [22]. We therefore first tested whether cDC2 were at all required for the induction of Th17 in our CD4⁺ T cell transfer model, in which we adoptively transferred Ovalbumin‐recognising CD4 T (OT‐2) cells into recipient mice, followed by intraperitoneal immunisation with OVA + LPS + αCD40, an established model to induce antigen‐specific CD4⁺ T cell priming in the mLNs. In accordance with previous studies using mice that lacked IRF4 in CD11c‐expressing cells resulting in a relative deficiency in cDC2 [11], huCD207.DTA mice lacking intestinal CD103⁺CD11b⁺ cDC2 showed significantly diminished potential of inducing Th17, showing that cDC2 were required for optimal Th17 induction in this model (Figure 5A). Interestingly, however, huCD207.MyD88^ON mice, in which only CD103⁺CD11b⁺ cDC2 express MyD88, were unable to support Th17 induction in the same model (Figure 5B). This suggests that while required, cDC2 were not sufficient, and cells other than cDC2 need to sense LPS for the optimal support of Th17 immunity.

FIGURE 5.

cDC2 are required for Th17 cell accumulation but are not sufficient to drive Th17 responses in an OT‐II transfer model. (A) CD207‐DTA or (B) human‐Langerin‐cre.MyD88^LSL/LSL [CD207.MyD88^ON], MyD88^LSL/LSL [KO] littermates and C57Bl/6 unrelated WT [control] mice were transferred with 1 × 10⁶ naïve CD4⁺ T cells isolated from CD45.1⁺ OT‐II mice. Twenty‐four hours later, mice were intraperitoneally injected with 0.5 mg ovalbumin, 25 μg anti‐CD40 and 20 μg LPS. FTY720 (20 μg) was administered on days 1 and 3 post‐immunisation. Mice were sacrificed 4 days after immunisation, and mesenteric lymph nodes were analysed for Th17 cells (CD45.1⁺TCRβ⁺CD4⁺CD62L⁻CD44⁺IL‐17A⁺). Each symbol represents one mouse. CD207‐DTA data (A) are pooled from 3 independent experiments (2 mice/group) and CD207.cre‐mediated MyD88 switch‐on experiments were performed twice with 3–5 mice/group. Statistical significance was determined using the Mann–Whitney U test (Figure 5A) or one‐way ANOVA with Tukey's post hoc (Figure 5B) (*p < 0.05, **p < 0.005, ***p < 0.0005, ****p < 0.0001).

4. Discussion

Sensing of pathogen associated molecular patterns by pattern recognition receptors such as TLRs on DCs initiates the cascade of DC migration and activation that is essential for the formation of adaptive immunity. Numerous studies assessed the importance of MyD88 in maintaining homeostasis and mounting immune responses by integrating signals downstream of most TLRs and the IL1R in the intestines and other organs [46, 47]. We here show that DC‐intrinsic MyD88 expression is sufficient for DC migration and that the rules governing DC migration in response to TLR engagement are broadly conserved across ages.

While the peculiarities of early life immune development and maturation are to date poorly understood, there is an appreciation that neonatal immunology differs in many aspects from that in adults. This is primarily attributed to specific needs at early age caused by the requirements to learn to tolerate the wealth of environmental antigens, while maintaining a protective immune barrier to pathogens in the absence of immune memory. A better understanding of early life immunity is, however, needed, as epidemiological and experimental data cement the notion that early life adverse events can lead to irreversible immune dysfunction later in life, while the world remains in dire need of efficacious vaccines for safe use in newborns. DCs are the main immune sentinels of the body and essential drivers of adaptive immunity. We here show that intestinal DCs are present in reduced numbers already shortly after birth and that TLR stimulation induces their migration to the draining mLN in a similar manner as in adults. While we have not tested the capacity of neonatal DCs to produce cytokines and induce adaptive immunity in our study, their similar migration expands on previous work showing that splenic neonatal DCs can efficiently process antigen and activate T cells [28]. Likewise, while neonatal cDC2 were previously shown to be under‐represented in the Peyer's Patch (but not the small‐intestinal lamina propria), they could be activated using R848 [40]. Altogether, this suggests that neonatal cDC1 and cDC2 might not be intrinsically different, but rather that prominent immune regulation during early life ensuring immune homeostasis may be caused by the different environment, including additional subsets of antigen‐presenting cells over‐represented at early age [48]. For example, several recent studies described RORγt⁺ APCs as non‐redundant tolerance‐inducing accessory cells over‐represented in peri‐weaning mice with the ability to process and present antigen in immune inductive sites [48, 49, 50].

We have shown previously that the TLR3/TRIF pathway is essential for DC activation and migration in response to poly(I:C), but that this requirement was not cell intrinsic, as cDC2, not expressing TLR3, migrated as efficiently as cDC1 [32]. Others have shown that pDC‐derived TNFα and type I IFN were required for the migration of intestinal cDC in response to oral feeding of the TLR7‐ligand R848, but whether MyD88 expression was required in the migrating cells was not assessed. Here, we found that DC migration in response to R848 was significantly blunted if only CD103⁺CD11b⁺ cDC2 expressed MyD88 but comparable to wildtype levels in the presence of MyD88 on all CD11c⁺ cells (including pDCs and macrophages). pDCs respond vigorously to TLR7 stimulation, which has not been shown for intestinal cDC2. It thus appears that, in response to R848, cDC rely on pDC‐mediated activation and instruction to migrate to the draining lymphoid tissues, in line with what was previously described [51]. In contrast, cDC2 express high levels of TLR5 [43], which we here show to lead to efficient FliC‐induced DC migration in the absence of MyD88 on any other cell subset. Together, MyD88 expression in cDC2 seems sufficient to induce migration if the sensing TLR is equipped by cDC2. A limitation of our study is that we have not formally excluded TLR7 expression and signalling in cDC2 from huCD207.MyD88 ^ON mice. Interestingly, some neonatal cDC1 maintained migration in response to R848 treatment in CD11c.MyD88 ^OFF mice, a finding that we currently cannot explain, but that might be caused by different environmental cues, by the availability of CD11c‐negative cells in the neonatal intestine that respond to TLR7, or different expression patterns of TLRs across ages. Effects might also be due to non‐TLR agonists signalling through MyD88 on bystander cells, such as IL‐33, which was previously shown to exert a pronounced effect on DC function, especially in the neonatal lung [47]. One caveat of our study is that we did not label intestinal DCs to track their migration to the LN, and it seems possible that R848 induces premature LN enlargement that would facilitate additional DC entry directly from the blood.

MyD88 expression in either epithelial cells or CD11c⁺ mononuclear phagocytes is sufficient to control acute intestinal infection with Clostridium difficile . This control is entirely independent of DC‐induced adaptive immunity and relies on innate defence mechanisms at the intestinal wall [52]. In response to Citrobacter rodentium , selective expression of MyD88 in all CD11c⁺ mononuclear phagocytes is sufficient for the induction of colonic adaptive Th1/Th17 responses [53]. The need for bona fide DCs for the induction of Th17 was recently challenged [22, 54], and CD11c and/or zbtb46, promoters that were used ubiquitously in genetic models to claim cDC function, were shown to be expressed by additional accessory cells of the intestines, including macrophages, innate lymphoid cells and other recently characterised antigen‐presenting cells, including Thetis cells and Janus cells, whose relationship to cDCs is under debate [22, 48, 49, 55, 56]. We here report that, as previously found in steady state in the intestinal lamina propria [42], the induction of Th17 in the mLNs in a T cell transfer model was significantly blunted in the absence of cDC2. Th17 induction, however, also did not occur if MyD88 expression was restricted to cDC2, suggesting that cells other than cDC2 must be involved in the Th17‐inducting cascade in this model.

In summary, we have shown that neonatal intestinal DC are present in early age and readily migrate to immune inductive sites upon immune activation, following similar kinetics and molecular requirements as their adult counterparts. This work provides essential fundamental knowledge for the design of efficacious oral vaccines for use in early life.

Author Contributions

K.G.M., S.H., A.G.L., C.S. and K.L. designed the experiments. K.G.M., S.H., A.G.L., S.C., C.S., I.U., J.N., J.H. and K.L. performed the experiments. K.G.M., S.H., A.G.L., C.S., J.H. and K.L. analysed the data. A.F.C. provided reagents and expertise. K.G.M. and K.L. supervised the work. K.L. wrote the paper. K.L. obtained funding. All authors reviewed and approved the final manuscript.

Funding

This work was supported by Ragnar Söderbergs stiftelse, Canadian Institutes of Health Research, GLB – 192248, Vetenskapsrådet, 2021‐01385.

Ethics Statement

All animal experiments were performed in accordance with European regulation and federal law of Denmark and Sweden and approved by the Danish Animal Experiments Inspectorate or the Lund/Malmö Animal Ethics Committee. Experiments at the University of Calgary were performed following the guidelines of the Canadian Council for Animal Care and approved by the University of Calgary Health Science Animal Care Committee (protocol #AC23‐0172).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Gating strategy for Figure 1.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.