Research ArticleINNATE IMMUNITY

Peripheral lymph nodes contain migratory and resident innate lymphoid cell populations

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Science Immunology  31 May 2019:
Vol. 4, Issue 35, eaau8082
DOI: 10.1126/sciimmunol.aau8082

Innate drifters

Innate lymphoid cells (ILCs) are regarded as tissue-resident sentries that stay put in peripheral tissues or lymphoid organs rather than extensively recirculating like most B and T cells. Dutton et al. used direct illumination to mark all cells in a single peripheral lymph node of transgenic mice expressing a photoconvertible fluorescent reporter protein. Migration versus continuing residence of the marked cells was monitored by flow cytometry. This experimental approach showed that ILC1s in a lymph node have more propensity to migrate away from that lymph node than ILC2s and ILC3s but still considerably less than T cells. The findings from this study reveal that some ILC subsets are not exclusively sessile and routinely traffic between different anatomic sites while carrying out their immunosurveillance mission.

Abstract

Tissue residency is considered a defining feature of the innate lymphoid cell (ILC) populations located within mucosal and adipose tissues. ILCs are also present within all lymphoid tissues, but whether ILCs migrate between lymphoid and nonlymphoid sites and in what context is poorly understood. To determine whether migratory ILCs exist within peripheral lymph nodes (LNs), we labeled all cells within the brachial LN (bLN) of transgenic mice expressing a photoconvertible fluorescent protein by direct exposure to light. Tracking of cellular changes in the labeled LN revealed the gradual migration of new ILCs into the tissue, balanced by egress of ILCs dependent on sphingosine-1-phosphate receptors. Most of the migratory ILCs were ILC1s, entering LNs directly from the circulation in a CD62L- and CCR7-dependent manner and thus behaving like conventional natural killer (cNK) cells. Upon egress, both ILC1s and cNK cells were found to recirculate through peripheral LNs. A distinct population of migratory ILC2s were detected in the LN, but most of the ILC3s were tissue resident. Functionally, both migratory and resident ILC1s within LNs were able to rapidly produce IFN-γ to support the generation of robust TH1 T cell responses after immunization. Thus, migratory and resident ILC populations exist within peripheral LNs, with ILC1s, akin to cNK cells, able to traffic into these tissues where they can contribute to the initiation of adaptive immunity.

INTRODUCTION

Innate lymphoid cells (ILCs) have been identified in most tissues in the body where they act as sentinels, orchestrating homeostasis, but able to dynamically respond to a wide range of signals indicative of infection or danger (13). As ILC populations were discovered, it became evident that these cells were most numerous at barrier tissues such as the gastrointestinal tract and lung, and the majority of studies have focused on their roles at these sites (48). To address whether ILCs were tissue resident, several studies have used parabiosis models, which indicated that the vast majority of ILCs at these barrier sites, as well as in adipose tissue, permanently reside within these tissues and do not traffic between host mice when the circulatory system is shared (912). Current models of ILC development propose the seeding of tissues with progenitors that develop from both fetal liver and bone marrow and then expand to provide tissue-resident populations (1317).

Although less numerous than at mucosal sites, ILCs are also present within all lymphoid tissues, suggesting roles influencing the adaptive immune responses initiated in these structures (18, 19). Crucially, how ILC populations within lymphoid tissue relate to those within barrier tissues is poorly understood. Analysis of the mesenteric lymph node (mLN) and spleen of parabiotic mice also suggested that most ILCs within these secondary lymphoid tissues were tissue resident (9). Furthermore, given the role of the lymphoid tissue–inducer subset of ILC3s in LN and Peyer’s patch organogenesis (20), it is possible that such LN-resident populations may be established very early in life. To date, the lack of mouse models that specifically and completely delete ILC populations has hindered our ability to define their functions within lymphoid tissues (21). Beyond their ability to rapidly produce effector cytokines, investigations of how ILCs might regulate adaptive immune responses have identified that both ILC2s (22) and ILC3s (19, 23) are able to process and present peptides via major histocompatibility complex II (MHCII) and modulate CD4 T cell responses. Where and how ILCs might acquire such antigen is unresolved but relevant to understanding the role such antigen presentation plays within the immune system. It is conceivable that, as a consequence of antigen uptake, ILCs migrate from tissue to draining LNs via the lymphatics. Such trafficking would not be evident within the parabiosis studies used to identify tissue residency. In support of ILC migration from tissues to their draining LNs, direct trafficking of ILCs from the intestine to the mLN has been described (18), and analysis of the mLNs of mixed bone marrow chimeras using wild-type (WT) and CCR7−/− donors indicated a cell-intrinsic requirement for CCR7 in establishing normal ILC populations within LNs (18). Because many studies of human ILCs rely on isolating these cells from blood, this suggests that circulatory ILCs exist at least in human blood; however, recent data have indicated that the human ILCs isolated from blood include progenitor populations (16, 24, 25). In addition, with better understanding of ILC subsets, a highly migratory ILC2 population was recently described (26), suggesting that the tissue-resident description of ILCs may not be true in all subsets in all circumstances. Last, conventional natural killer (cNK) cells, which share many similarities with ILC1s, are thought to be highly migratory, entering LNs directly from the circulation via high endothelial venules (HEV) (27). Thus, the dogma of ILC tissue residency needs to be further tested for lymphoid ILC populations. As we seek to better understand the roles ILCs play in secondary lymphoid tissue, how and when ILCs establish residency within LNs will help to clarify their role in the generation and regulation of adaptive immunity.

To test the hypothesis that ILCs in peripheral LNs are migratory, we have used Kaede photoconvertible mice (28), having established methods to label all cells within a peripheral LN by illumination after surgical exposure (29). This approach offers direct in vivo assessment of the migration of new cells into the tissue, identification of resident populations, and tracking of cells leaving the LN to new locations. Our data reveal that LNs contain both migratory and resident ILC populations, with the ILC1 subset highly migratory and able to recirculate through peripheral lymphoid tissue, where it can contribute to supporting the priming of CD4+ T cells after immunization.

RESULTS

ILCs migrate into peripheral LNs

To test the hypothesis that migratory ILCs exist in peripheral LNs, we exposed the brachial LN (bLN) of Kaede mice through a small incision in the skin and photoconverted as described previously, changing the cellular fluorescence from Kaede Green to Kaede Red (29). Although cells expressing the Kaede Red form of the fluorochrome frequently also express the Kaede Green form (due to a combination of incomplete conversion of total Kaede protein within the cell and the generation of new protein after photoconversion), to simplify the description of the data, here these cells are termed “Kaede Red+”. Analysis of Kaede expression confirmed that the entire hematopoietic compartment of the LN was converted to Kaede Red+ cells (fig. S1). Thus, analysis of the labeled bLN at subsequent time points after photoconversion allows identification of cells newly entering the tissue from unexposed sites (Kaede Green+) and those that have remained within the LN (Kaede Red+). A fraction of cells identified as Kaede Red+–resident cells may be migratory cells that have recirculated back to the initially photoconverted LN. This experimental approach enables the simple but direct assessment of residency within the LN for any cell type. After 72 hours, about 95% of the CD45+ cells in the labeled LN were Kaede Green+ cells (Fig. 1, A and B), consistent with the constant circulation of naïve lymphocytes through secondary lymphoid tissue. The majority of the Kaede Green+ cells within LNs were αβ T cells, although some γδ T cells were also detected (Fig. 1, C and D). A small but distinct population of IL-7Rα+Lin (B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, and FcεRI) ILCs was evident and comparable in frequency with Langerhans cells (LCs; Fig. 1, C and D). These data indicate that some ILCs had entered the tissue subsequent to photoconversion of the LN 72 hours previously. Among the Kaede Red+ cells that had remained within the LN since labeling, ILCs were also detected and appeared enriched compared with the proportion of ILCs among the Kaede Green+ population (Fig. 1, E and F). Thus, our initial analysis suggested the presence of both migratory and resident ILC populations within peripheral LNs.

Fig. 1 CCR7-dependent entry of ILCs into peripheral LNs.

To assess the migratory kinetics of different cellular populations within peripheral LNs, the bLN of Kaede mice was photoconverted and analyzed 72 hours later. (A) Expression of Kaede Green (KG) versus Kaede Red (KR) protein by CD45+ cells isolated from the bLN at 72 hours after photoconversion. (B) Percentage of CD45+ cells expressing Kaede Green or Kaede Red protein at 72 hours after photoconversion (n = 10). (C) Characterization of Kaede Green+ CD45+ cells isolated from the bLN, showing αβ T cells (CD45+ CD3+ TCRβ+ TCRγδ), γδ T cells (CD45+ CD3+ TCRβ TCRγδ+), ILCs (CD45+ IL-7Rα+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}), and LCs (CD45+ MHCII+ CD11c+ CD207+). (D) Percentage of αβ T cells (n = 7), γδ T cells (n = 7), ILCs (n = 10), or LCs (n = 5) among CD45+ Kaede Green+ cells. (E) Characterization of Kaede Red+ cells isolated from the bLN, showing αβ T cells, γδ T cells, ILCs, and LCs. (F) Percentage of αβ T cells (n = 7), γδ T cells (n = 7), ILCs (n = 10), and LCs (n = 5) among CD45+ Kaede Red+ cells. Expression of Kaede Green versus Kaede Red protein by αβ T cells, γδ T cells, ILCs, and LCs isolated from the bLN of (G) Kaede and (H) CCR7−/− Kaede mice. (I) Number of αβ T cells (WT, n = 7; CCR7−/−, n = 8), γδ T cells (WT, n = 7; CCR7−/−, n = 8), ILCs (WT, n = 6; CCR7−/−, n = 8), and LCs (WT, = 5; CCR7−/−, n = 7) isolated from the bLN of WT Kaede versus CCR7−/− Kaede mice. (J) Number of Kaede Green αβ T cells (WT, n = 7; CCR7−/−, n = 8), γδ T cells (WT, n = 7; CCR7−/−, n = 8), ILCs (WT, n = 7; CCR7−/−, n = 8), and LCs (WT, n = 5; CCR7−/−, n = 7) isolated from the bLN. Data are pooled from a minimum of two independent experiments. Values on flow cytometry plots represent percentages; bars on scatter plots represent the median, which is also shown numerically. Statistical significance was tested using an unpaired, nonparametric, Mann-Whitney two-tailed U test: *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001.

Although the majority of the CD45+ population in LNs are naïve lymphocytes and thus expected to be highly migratory, the CD45 stromal compartment would be expected to be obligatorily tissue resident. Therefore, we assessed Kaede expression in LN stromal cells to further confirm that our experimental model properly distinguished between resident and migratory populations. Blood endothelial cells, fibroblastic reticular cells, and lymphatic endothelial cells were all found to remain >96% Kaede Red+ at 72 hours after photoconversion (fig. S1), consistent with being resident populations.

To specifically assess the relative migratory abilities of individual cell types, we first identified different immune cell populations, and then, we analyzed Kaede Green versus Kaede Red expression. These data showed that, for αβ and γδ T cells, the vast majority (>93%) had trafficked into the LN within the preceding 72 hours (Fig. 1G). Unexpectedly, analysis of the ILC compartment revealed that about half the population had newly arrived into the tissue within the 72-hour time frame, arguing that a substantial proportion of ILCs in the LN were not tissue resident (Fig. 1G). To better understand the kinetics of ILC trafficking into LNs, we assessed T cell and ILC populations at different times after photoconversion (fig. S2). Although more than 90% of the LN T cell compartment was Kaede Green+ 24 hours after photoconversion, only 20% of the ILCs were Kaede Green+, suggesting that the rate of replacement of LN ILCs within peripheral LNs with newly migrating cells was substantially slower than for naïve T cells. Given the importance of CCR7 in the recruitment of cells into LNs, either via the lymphatics or from the blood (30), the bLN of CCR7−/− Kaede mice was assessed at 72 hours after photoconversion (Fig. 1H). For both T cells and ILCs, the proportion of Kaede Red+ cells increased compared with WT Kaede controls, however, this reflected the significantly reduced cellularity of the CCR7−/− tissue due to the inability of cells to enter and exit many tissues in the body (Fig. 1I). Nonetheless, consistent with a requirement for CCR7 for T cell and ILC entry into LNs, the number of Kaede Green+ T cells or ILCs was substantially reduced (Fig. 1J). Combined, these data reveal that both migratory and resident ILC populations exist within peripheral LNs, and the entry of migratory ILCs into these tissues is CCR7 dependent.

ILCs egress from LNs in an S1P-dependent manner

Because labeling of the LN was performed after a small excision in the skin, it was possible that the Kaede Green+ ILCs observed in the LN reflected additional migration of ILCs in response to local tissue damage, rather than normal homeostatic trafficking of these cells into the tissue. Enumeration of the number of Kaede Green+ versus Kaede Red+ ILCs in control mice (no photoconversion) compared with 72 hours after photoconversion indicated that the total number of ILCs in the LN was not increased by the process of labeling (Fig. 2, A and B), suggesting that new cells had not migrated into the tissue. Furthermore, quantitation of ILC numbers in the bLN of WT mice that had never undergone surgery versus those that had experienced a skin incision and photoconversion 72 hours previously revealed no differences in ILC numbers (fig. S3).

Fig. 2 ILC frequency in LNs is maintained by balanced ingress and egress.

To further investigate the migratory ILCs identified within the labeled bLN, the effect of photoconversion on the number of ILCs was assessed. (A) Expression of Kaede Green versus Kaede Red protein by ILCs (CD45+ IL-7Rα+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}) isolated from the bLN of control Kaede mice (no photoconversion) and Kaede mice at 72 hours after photoconversion. (B) The number of Kaede Green+ and Kaede Red+ ILCs recovered from the bLN of control (n = 6) versus photoconverted bLNs at 72 hours after labeling (n = 10). To investigate egress of ILCs from the bLN, Kaede mice were maintained on vehicle control (H2O) or FTY720, and the bLN was photoconverted and then analyzed 72 hours later. (C) Representative flow cytometry plots showing expression of Kaede Green versus Kaede Red protein by ILCs in the bLN 72 hours after photoconversion. (D) Percentage of Kaede Green+ or Kaede Red+ ILCs in the bLN of control (Ctrl; n = 4) and FTY720 (n = 7) mice. (E) Total numbers of Kaede Green+ and Kaede Red+ ILCs in the bLN. To assess whether ILCs recirculate through peripheral LNs, the contralateral LNs of Kaede mice were analyzed at 72 hours after photoconversion of the bLN, with combined B and T cell depletion used to enrich ILCs. (F) Expression of Kaede Green versus Kaede Red protein by ILCs (CD45+ IL-7Rα+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}) isolated from contralateral LNs. Percentage of Kaede Red+ ILCs (G) and T cells (H) in WT Kaede (n = 14) and CCR7−/− Kaede (n = 8) contralateral LNs. (I) Representative flow cytometry plots of Kaede Green+ (left) and Kaede Red+ (right) ILC expression of CD69. (J) Percentage of Kaede Green+ and Kaede Red+ ILCs expressing CD69 (n = 7). Data are pooled from two independent experiments. Bars on scatter plots represent the median, which is also shown numerically. Statistical significance was tested using an unpaired, nonparametric, Mann-Whitney two-tailed U test: **P ≤ 0.01, ***P ≤ 0.001.

These data indicate that ILC migration into the tissue was matched by the loss of ILCs either through cell death or egress from the LN. The egress of B and T cells from LNs is dependent on the sensing of sphingosine-1-phosphate (S1P) by S1P receptor 1 (S1P1) (31), whereas the egress of cNK cells from LNs is dependent on S1P5 (32, 33). Expression of S1P1 was detected on a proportion of ILC1s, ILC2s, and ILC3s isolated from the bLN (fig. S4), suggesting that ILCs might use S1P-S1P1 interactions to mediate egress from peripheral LNs, as described for T cells. To test whether ILCs egress from LNs, we performed photoconversion of the bLN on mice maintained on vehicle control (H2O) or the S1P receptor agonist FTY720, a small-molecule inhibitor that blocks lymphocyte recirculation through inhibiting three of the five S1P receptors: S1P1, S1P3, and S1P4 (31, 34). The total number of T cells within the LN was not significantly altered by FTY720 treatment (fig. S5); however, analysis of Kaede Red expression confirmed that this was due to the expected retention of T cells within the LNs (fig. S5), which, in turn, meant that far fewer Kaede Green+ T cells were migrating into the tissue (fig. S5). Analysis of ILCs revealed an increase in the proportion of Kaede Red+ ILCs (Fig. 2, C and D) and a significant increase in the total number of Kaede Red+ ILCs in the LN, whereas the number of Kaede Green+ ILCs remained unchanged through FTY720 treatment (Fig. 2E). Thus, inhibition of S1P receptor–mediated exit from the tissue resulted in the accumulation of ILCs within the LN, arguing that under normal conditions, these cells egress from the tissue. To further investigate expression of S1P receptors by ILCs, we performed gene expression analysis on sorted ILC1 (NK1.1+NKp46+) and non-ILC1 (NK1.1NKp46) ILC populations isolated from peripheral LNs by fluorescence-activated cell sorting (FACS). S1P1 and S1P4, both of which are inhibited by FTY720, were observed to be most highly expressed by LN ILC populations, whereas relatively little expression of S1P5 was detected (fig. S6). The sorted ILC1 population, which expressed significantly higher levels of interferon-γ (IFN-γ) and T-bet than the other LN ILCs, also expressed significantly higher levels of CD62L, required for entry into LNs from HEV (fig. S6).

Because ILCs that have left the photoconverted LN can be identified as Kaede Red+, the ability of ILCs to recirculate through peripheral LNs was tested by assessing Kaede Red+ ILCs in a pool of contralateral LNs. ILCs were enriched (through depletion of B and T cells) to aid the identification of rare Kaede Red+ ILCs, and this approach could detect a small Kaede Red+ ILC population that constituted only a very small proportion of the ILCs isolated from the contralateral LNs (0 to 0.7% of the population; Fig. 2, F and G), although this was more than detected in the contralateral LNs of CCR7−/− Kaede mice. This low frequency of recirculating ILCs may represent either the limitations of tracking rare populations, alongside some ILCs migrating to tissues other than LNs. T cells recirculating from the photoconverted LN comprised about 2.5% of the T cell population in contralateral LNs (Fig. 2H and fig. S5). To further assess expression of molecules associated with retention of cells within tissues, we assessed expression of CD69 on Kaede Green+ and Kaede Red+ ILCs. Kaede Green+ ILCs expressed very little CD69, whereas most of the Kaede Red+ ILCs were CD69+, consistent with a tissue-resident phenotype (Fig. 2, I and J). In summary, these data show that the migration of ILCs into peripheral LNs is accompanied by the egress of these cells in an S1P receptor–dependent manner, and at least some of these ILCs then recirculate through other peripheral LNs.

ILC1s are the migratory ILC subset

To determine whether all ILC populations were equally migratory, we sought to define the Kaede Green+ and Kaede Red+ ILCs within the bLN. ILC subsets in the bLN can be readily distinguished by staining for transcription factors (TFs; fig. S7) (18). T-bet+ ILC1s in the bLN lacked expression of eomesodermin (EOMES) (i.e., T-bet+EOMES), demonstrating that the ILC1s we had identified were not misidentified cNK cells (fig. S7). In photoconverted LNs, methods to preserve Kaede protein expression resulted in suboptimal TF detection. Therefore, a panel of surface markers that identified the ILC subsets in the bLN of WT mice was devised (fig. S7) and used to identify different ILC subsets only by surface markers (Fig. 3, A and B). Analysis of Kaede Green versus Kaede Red protein expression in bLN ILCs at 72 hours after photoconversion revealed that there were relatively few migratory ILC2s and ILC3s (~20% Kaede Green+), whereas ILC1s were much more migratory because more than 60% were Kaede Green+ (Fig. 3C). Similar proportions of Kaede Green+ ILCs were also observed at 1 week after photoconversion (Fig. 3D), indicating that LNs contained both migratory and resident ILC populations, with different subsets showing distinct migratory properties (Fig. 3E). ILC tissue residency within the LN was then further assessed using a second photoconvertible mouse strain, the H2B-Dendra2 model (35). The fusion of the Dendra protein to histones enabled intracellular staining of TFs to be used to more definitively identify ILC subsets. In H2B-Dendra2 mice, ILC subsets could be split into resident and migratory populations in a comparable manner with that observed in the Kaede mice (fig. S8). Identification of ILC2s based upon GATA-3 expression, rather than surface markers used in Kaede mice, indicated a larger migratory ILC2 population within the LN (fig. S9). Because the expression of Kaede Red protein is lost over several days in rapidly proliferating cells due to label dilution, the emergence of Kaede Green+ ILCs could theoretically be explained by highly proliferative populations in the tissue (28). Analysis of Ki-67 expression within the bLN revealed that most of each ILC subset was Ki-67, and the proportions of Ki-67+ ILC1s and ILC2s were comparable, despite very different patterns of Kaede Red and Kaede Green expression at 72 hours after photoconversion (fig. S10). Thus, the high proportion of Kaede Green+ ILC1s cannot be explained by Kaede Red protein dilution through proliferation.

Fig. 3 ILC1s traffic through peripheral LNs from the circulation.

To test whether all ILC subsets showed an equal ability to traffic through the peripheral LNs, an established panel of surface markers was used to identify which ILC subsets were migrating into the bLN. (A) Representative flow cytometry plots showing identification of ILC1 (CD45+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI} IL-7Rα+ CD4 (NK1.1, NKp46)+), ILC2 (CD45+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}) IL-7Rα+ CD4 (NK1.1, NKp46)(KLRG-1, CD25)+), and ILC3 (CD45+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}IL-7Rα+ CD4+] subsets in the bLN. (B) The percentage of each ILC subset identified within the bLN of Kaede mice using the surface marker staining. (C and D) Kaede Green versus Kaede Red protein expression by ILC1s, ILC2s, and ILC3s at 72 hours and 1 week after photoconversion of the bLN. (E) The percentage of Kaede Green+ and Kaede Red+ ILC1s, ILC2s, and ILC3s in the bLN at 72 hours and 1 week after photoconversion. Data are pooled from two independent experiments, n = 7. To investigate the source of ILC1s migrating into the bLN, ILC populations in murine blood were analyzed. (F) Representative flow cytometry plots showing identification of putative ILC1s (CD45+ IL-7Rα+ (NKp46NK1.1)+Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}) and ILC2s (CD45+ IL-7Rα+ NK1.1 NKp46 KLRG-1+ Lin). (G) The proportion of ILC1s and ILC2s within the ILC population isolated from the blood, n = 8. (H) Representative flow cytometry plots showing expression of T-bet versus GATA-3 by blood ILCs. To isolate sufficient numbers of ILCs, blood samples from three mice were enriched for the CD45+ fraction separately and then pooled for staining and analysis. (I) Representative flow cytometry plots showing expression of CD62L and CCR7 by T-bet+ ILC1s versus T-bet ILCs isolated from blood. All data are pooled from two independent experiments. Values on flow cytometry plots represent percentages; bars on scatter plots represent the median. Statistical significance was tested using an unpaired, nonparametric, Mann-Whitney two-tailed U test: ***P ≤ 0.001.

The IL-7Rα+T-bet+EOMES ILC1s assessed within the LN are developmentally distinct from cNK cells, despite both being grouped in the ILC1 subset and cNK cells having been previously shown to enter LNs direct from the circulation (27). To directly compare ILC1 and cNK cell migration, we identified these two populations within the photoconverted bLN at 72 hours after labeling and in a pool of contralateral LNs (fig. S11). Although ILC1s were 60% Kaede Green+, cNK cells were 90% Kaede Green+, indicating that, whereas the LN compartment of ILC1s contains both migratory and resident populations, the vast majority of cNK cells are highly migratory. Analysis of contralateral LNs identified a small Kaede Red+ ILC1 population and a larger Kaede Red+ cNK cell population, demonstrating that both populations recirculate through peripheral LNs.

Given that cNK cells are reported to enter LNs via HEV (27), ILC populations in mouse blood were assessed to investigate whether ILC1s in the murine circulation might be the source of the ILCs trafficking into the bLN. A very small (<0.05 to 0.011% total CD45+ cells) IL-7Rα+Lin population was detected in the circulation, the majority of which expressed both NK1.1 and NKp46 at steady state alongside a few putative ILC2s (NK1.1KLRG-1+) (Fig. 3, F and G). These ILCs also expressed T-bet (Fig. 3H), providing further evidence of an ILC1 phenotype and consistent with previous data suggesting that ILC1s were present in the circulation (9, 11). To investigate whether ILC1s in the blood were equipped to enter LNs directly from the circulation via HEV, we assessed the expression of CD62L and CCR7. About half of blood ILC1s expressed both CD62L and CCR7, whereas the few other ILCs (IL-7Rα+Lin cells) detectable in blood appeared to lack expression of these molecules (Fig. 3I). Combined, our analysis of ILC subsets revealed that ILC1s show behavioral similarities to cNK cells, and a circulating ILC1 population in the blood is able to traffic into and out of peripheral LNs.

ILCs do not migrate from the skin to peripheral LNs under steady-state conditions

Although our data were consistent with ILC migration into LNs arising from the circulation, it was possible that ILCs also trafficked from skin (36) to peripheral LNs via the lymphatics, as observed for dermal dendritic cells (DCs) and LCs (37). To directly test this, we established photoconversion of the entire CD45+ compartment of ear skin, enabling the analysis of cellular migration into the draining auricular LN (auLN) and identification of new cells moving into the ear skin (Fig. 4, A and B). To directly test the migration of cells from the ear skin to the auLN, we photoconverted the ear and analyzed the auLN 48 and 72 hours later. A modest, but distinct, Kaede Red+ population was detected within the auLN (Fig. 4, C to E), with T cells, but no ILCs were detected among this small migratory population (Fig. 4, F and G). These data argue that, under steady-state conditions, ILCs do not migrate from skin to peripheral LNs. To determine whether ILCs in skin only migrate to peripheral LNs during inflammation, the MC903-induced atopic dermatitis model (38) was established in Kaede mice, combining 5 days of MC903 ear painting with photoconversion of the skin. The short 5-day treatment was chosen to enable early changes in ILC populations to be assessed and caused a substantial increase in the size of the auLN, significant increases in ear thickness, and increased numbers of basophils and other hematopoietic cells in the ear skin including ILC2s (fig. S12) (39). The draining auLN had increased substantially in cellularity, including the number of ILCs, which had increased about 15-fold (Fig. 4, H to J). Analysis of Kaede Red+ cells in the auLN revealed increased numbers of cells trafficking to the draining LN (Fig. 4, K and L), which were not only mostly T cells but also ILCs (Fig. 4, M and N). Detection of such few migratory ILCs compared with the substantial increases in all ILC subsets observed in the auLN (Fig. 4O) argued that migration through the lymphatics was unlikely to be the main mechanism increasing ILC numbers in the LN. ILCs had significantly increased their expression of Ki-67 (Fig. 4P), suggesting at least that the increased numbers of ILCs in the auLN reflect local expansion in situ.

Fig. 4 Minimal ILC migration from skin to the draining LN even under conditions of inflammation.

To directly test whether ILCs migrate from skin to peripheral LNs, the ears of Kaede mice were photoconverted, and the presence of labeled cells were assessed in the auLN. (A) Expression of Kaede Green versus Kaede Red protein by CD45+ cells isolated from ear skin without (left) or immediately after (right) photoconversion. (B) Percentage of CD45+ cells expressing Kaede Red protein in the ear and auLN immediately after photoconversion (n = 12). (C) Expression of Kaede Green versus Kaede Red protein by CD45+ cells in the auLN at 0, 48, and 72 hours after photoconversion of the ear skin. The percentage (D) and total number (E) of Kaede Red+ CD45+ cells in the auLN over the time course of 0 hours (n = 12), 48 hours (n = 11), and 72 hours (n = 10). (F) Representative flow cytometry plots assessing the phenotype of Kaede Red+ CD45+ cells in the auLN, with identification of T cells (CD3+), ILCs (IL-7Rα+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}), and LCs (CD45+ MHCII+ CD11c+ CD207+). (G) Percentage of Kaede Red+ CD45+ cells in the auLN that were T cells (n = 5), ILCs (n = 12), and LCs (n = 5). To determine whether inflammation in the skin resulted in the migration of ILCs to the draining LN, the ears of mice were treated with vehicle control (VC) or MC903, with photoconversion of the ear after 48 hours of treatment. (H) Representative flow cytometry plots showing ILCs within VC-treated (left) and MC903-treated (right) auLN. Enumeration of CD45+ cells (I) and ILCs (J) recovered from the auLN of mice treated with VC (n = 7) or MC903 (n = 12). (K) Representative flow cytometry plots assessing the phenotype of Kaede Red+ CD45+ cells in the auLN of MC903-treated mice, with the identification of T cells and ILCs. Enumeration of Kaede Red+ (L) CD45+ cells in VC (n = 8) or MC903 (n = 8), (M) T cells in VC (n = 10) or MC903 (n = 11), and (N) ILCs in VC (n = 8) or MC903 (n = 8) within the auLN. (O) Enumeration of ILC subsets including triple negative (TrN) in the auLN of mice treated with VC (n = 6) and MC903 (n = 15). (P) Expression of Ki-67 by ILCs in the auLN of mice treated with VC (n = 6) or MC903 (n = 7). Data are pooled from a minimum of two independent experiments. Values on flow cytometry plots represent percentages; bars on scatter plots represent the median, which is also shown numerically. Pairs of samples were compared using an unpaired Mann-Whitney U test. When comparing more than two sets of data, statistical significance was tested using Kruskal-Wallis one-way analysis of variance (ANOVA) with post hoc Dunn’s test: *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001.

Combined, these data reveal that, under steady state, ILCs traffic from the circulation into peripheral LNs rather than through lymphatic drainage from the skin. Although inflammation in the skin can cause some ILCs to migrate to the draining LN, the substantial increase in ILC numbers reflects other mechanisms such as local proliferation and recruitment from the blood.

Recruitment of ILCs into inflamed skin is dependent on CCR6

Simultaneous photolabeling of all the hematopoietic cells in the ear enabled migration of cells into the ear skin to be assessed. At 48 and 72 hours after photoconversion, the proportion of Kaede Green+ CD45+ cells in the ear had significantly increased compared with 0-hour controls, indicating the trafficking of new cells into this site at steady state (Fig. 5, A and B), of which ILCs were a small (~3%) proportion (Fig. 5, C and D). This may represent redistribution of ILCs within the skin or trafficking of these cells from the following education in the draining LN (40). To test whether the recruitment of ILCs into skin was enhanced during inflammation, we assessed Kaede Green+ cells migrating into the ear after administration of MC903. A significant increase in the number of Kaede Green+ CD45+ cells (Fig. 5E), T cells (Fig. 5F), and ILCs (Fig. 5G) was observed in MC903-treated mice versus vehicle control, suggesting that ILCs were recruited into the inflamed ear tissue. Over half of the population of Kaede Green+ ILCs lacked Ki-67 expression (Fig. 5H), arguing that the increase in Kaede Green+ ILCs was not solely attributable to proliferation. To investigate the potential mechanism regulating ILC recruitment to the inflamed tissue, we assessed CCR6 expression on ILCs given the proposed role of its ligand, CCL20, in regulating homing to the skin (41). Although expression of CCR6 is restricted to ILC3s in the mLN, all ILC subsets were isolated from ear skin or the auLN expressed CCR6, indicating that this might be a key mechanism by which these cells are recruited to the skin (fig. S13). To directly test this, we treated CCR6−/− Kaede mice and WT Kaede mice with MC903 as described above. Analysis of Kaede Green+ cells in the ear of CCR6−/− Kaede mice revealed a significant drop in the total number of CD45+ cells, which included a significant decrease in the number of ILCs (Fig. 5, I to L), indicating that CCR6 is important for the recruitment of ILCs into inflamed skin.

Fig. 5 CCR6-dependent recruitment of ILCs into the inflamed ear.

To investigate mechanisms controlling the recruitment of ILCs into the ear, the entire ear CD45+ population was labeled through photoconversion, with Kaede Green+ cells identified at subsequent time points as cells that had newly entered the tissue. (A) Expression of Kaede Green versus Kaede Red protein by CD45+ cells in the ear at 0, 48, and 72 hours after photoconversion of the ear skin. (B) The percentage (left) and total number (right) of Kaede Green+ CD45+ cells in the auLN over the time course of 0 hours (n = 12), 48 hours (n = 11), and 72 hours (n = 11). (C) Representative flow cytometry plots assessing the phenotype of Kaede Green+ CD45+ cells in the ear with identification of T cells (CD3+), ILCs (IL-7Rα+ Lin{B220, CD11b, CD11c, CD3, CD5, CD19, Ter119, Gr1, CD49b, F4/80, FcεRI}) and LCs (CD45+ MHCII+ CD11c+ CD207+). (D) Percentage of Kaede Green+ CD45+ cells in the ear that were T cells (n = 6), ILCs (n = 11), and LCs (n = 6). To assess migration under inflammatory conditions, MC903 was applied to the ears for five consecutive days to induce atopic dermatitis, with photoconversion of the ear 48 hours after treatment was started. Number of Kaede Green+ (E) CD45+ cells in VC (n = 9) or MC903 (n = 7), (F) T cells in VC (n = 10) or MC903 (n = 8), and (G) ILCs in VC (n = 9) or MC903 (n = 7) within the ear. (H) Representative flow cytometry plots of Kaede Green+ ILCs Ki-67 expression within the ear of VC- and MC903-treated mice. To test the requirement for CCR6 in ILC recruitment into the ear, MC903 was applied to the ears of Kaede and CCR6−/− Kaede mice for five consecutive days to induce atopic dermatitis, with photoconversion of the ear 48 hours after treatment was started. (I) Expression of Kaede Green and Kaede Red protein by CD45+ cells within MC903-treated ears. (J) Number of Kaede Green+ CD45+ cells (Kaede, n = 8; CCR6−/− Kaede, n = 9). (K) Representative flow cytometry plots showing Kaede Green+ ILCs. (L) Number of Kaede Green+ ILC (Kaede, n = 8; CCR6−/− Kaede, n = 9). Each data point represents cells isolated from one ear and one auLN. Data are pooled from a minimum of two independent experiments. Values on flow cytometry plots represent percentages; bars on scatter plots represent the median, which is also shown numerically. Pairs of samples were compared using an unpaired Mann-Whitney U test. When comparing more than two sets of data, statistical significance was tested using Kruskal-Wallis one-way ANOVA with post hoc Dunn’s test: *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001.

ILC1s are a rapid source of IFN-γ in the draining LN after immunization

Combined, our data revealed that a migratory ILC1 population was present within peripheral LNs, alongside a population that remained resident in the tissue for at least 1 week. The ability of cNK cells to rapidly produce effector cytokines is thought to modulate the microenvironment of the LN and shape the T cell responses generated in these tissues (42). Thus, we asked whether ILC1s in peripheral LNs also contributed to the rapid early release of cytokines in response to immunization. NK cell production of IFN-γ within the draining peripheral LNs was recently identified as a key mechanism through which the adjuvant AS01 drives the activation of DCs and the development of T helper 1 (TH1) immunity (43). To assess whether the ILC1 population in peripheral LNs can also contribute to this early IFN-γ response, we immunized WT mice intramuscularly with the antigen OVA-2W1S, in the presence of phosphate-buffered saline (PBS) or with AS01. Consistent with the previous study, AS01 induced the rapid production of IFN-γ by NK1.1+CD3 cells in the draining LN within hours of immunization (Fig. 6A). Although the majority of the NK1.1+CD3 cells expressed EOMES, a distinct population of NK1.1+CD3IL-7Rα+EOMES cells expressing T-bet were evident, thus revealing that ILC1s in the LN also release IFN-γ in response to AS01 (Fig. 6, A and B). Confirming that this response had resulted in a TH1-skewed T cell response, analysis of 2W1S-specific CD4 T cells in the draining LN at 7 days after immunization using MHCII tetramers revealed that the AS01 adjuvant had generated a robust TH1 effector T cell response (Fig. 6, C and D). Last, to determine whether the migratory and resident ILC1 populations showed functional differences in their production of IFN-γ, we immunized Kaede mice with OVA-2W1S in the presence of PBS or AS01, 72 hours after photoconversion of the draining bLN. Both Kaede Green+ and Kaede Red+ ILC1s in the draining LN expressed IFN-γ (Fig. 6E), although numerically, there were significantly more Kaede Green+ ILC1s among the IFN-γ–producing cells than Kaede Red+ ILC1s (Fig. 6F). The vast majority of IFN-γ produced by cNK cells came from Kaede Green+ cells (Fig. 6G). Therefore, both migratory and resident ILC1s appear to be able to functionally contribute to the shaping of adaptive immune responses within LNs through rapid effector cytokine release.

Fig. 6 ILC1s in the draining LN are a rapid source of IFN-γ after immunization.

To investigate the function of ILC1 within peripheral LNs, mice were injected intramuscularly with 20 μg of OVA-2W1S/PBS or 20 μg of OVA-2W1S/AS01 and culled either 6 hours after immunization to assess the early innate IFN-γ response or after 7 days after a second immunization performed 6 hours before analysis. (A) Representative flow cytometry plots showing inguinal LN cells from mice given OVA-2W1S/PBS (top) or OVA-2W1S/AS01 (bottom), with identification of different IFN-γ–producing populations including cNK cells (CD45+ IFN-γ+ CD3ε NK1.1+EOMES+) and ILC1s (CD45+ IFN-γ+ CD3ε NK1.1+ EOMES T-bet+). (B) Total number of IFN-γ+ ILC1s and cNK cells in OVA/ASO1-treated mice (n = 5). (C) Representative flow cytometry plots showing IFN-γ production by 2W1S-specific CD4+ T cells at day 7 after initial immunization with (AS01) or without (PBS) the adjuvant AS01. (D) Total number of IFN-γ producing 2W1S-specific CD4 T cells in the draining inguinal LN of PBS (n = 4) and AS01 mice (n = 6). To test whether both migratory and resident ILC1 subsets were able to produce IFN-γ, the bLN of Kaede mice was photoconverted, and 3 days later, mice were then immunized in the paw pad with either 20 μg of OVA-2W1S/PBS or 20 μg of OVA-2W1S/AS01 and analyzed 6 hours later. (E) Representative flow cytometry plots showing bLN cells from mice given OVA-2W1S/PBS (top) or OVA-2W1S/AS01 (bottom), with identification of ILC1 and cNK populations producing IFN-γ and their expression of Kaede Green versus Kaede Red protein. (F) Percentage Kaede Green (migratory) and Kaede Red (resident) ILC1s and cNK cells among IFN-γ–producing cells. (G) Total numbers of IFN-γ+ ILC1s and cNK cells expressing either Kaede Green (migratory) or Kaede Red (resident). All sets of data are pooled from two independent experiments. Values on flow cytometry plots represent percentages; bars on scatter plots represent the median, which is also shown numerically. Statistical significance was tested using an unpaired, nonparametric, Mann-Whitney two-tailed U test: *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001.

DISCUSSION

Here, we provide compelling evidence that peripheral LNs contain migratory ILC populations. Our studies revealed the balanced ingress and egress of ILCs from LNs with most of the migratory ILCs belonging to the ILC1 subset over the time frame assessed. ILC1s are the most prevalent subset in murine circulation and, like cNK cells, recirculate through peripheral LNs using CCR7 and CD62L to enter the tissue while egressing in an S1P receptor-dependent manner. Functionally, ILC1s in peripheral LNs were able to rapidly produce effector cytokines in response to immunization, as observed for cNK cells, supporting the initial priming of CD4 T cells. Combined, these data provide critical further insight into the behavior of ILCs within lymphoid tissues.

After their relatively recent identification (3), understanding the functions of ILCs within the immune system has become a key area of immunological research. Despite ILCs being present within all lymphoid tissues, direct evidence of their function here is very limited. Having established techniques to track the movement of cells into and out of peripheral LNs, we sought to understand more of the basic behavior of ILCs within these tissues in an effort to better understand their roles within such tissue. Photoconvertible mice provide a means of site specifically labeling cells and the direct assessment of cellular migration in vivo. Tracking ILCs within the photoconverted bLN over time revealed that, within 72 hours, about 50% of the ILCs had migrated into the tissue since photoconversion. Although the fluorescence level of Kaede Red protein will dilute as cells proliferate, most of the ILCs within the LN were Ki-67, and expression of Kaede Red did not correlate with proliferation among ILC subsets. Although a clear caveat of the photoconvertible mouse models is the potential loss of the light-induced label, studying ILC migration under steady-state conditions over a time frame of up to 1 week helped to mitigate this problem. Treatment with the S1P receptor agonist FTY720 resulted in the accumulation of ILCs within the LN, consistent with blocked egress. These data strongly indicated the balanced migration of ILCs into and out of peripheral LNs. Combined with our data detecting minimal ILC trafficking from skin to the draining peripheral LN, the detail of which ILC subsets migrate really reflects the composition of the ILC compartment within murine blood. Here, most of the ILCs were of an ILC1 phenotype, and expression of the key mechanisms required to enter LNs direct from the circulation (CD62L and CCR7) was detected. Only a minority of the ILC2 and ILC3 populations within the LN were found to be migratory over 1 week, indicating that bona fide LN-resident ILC populations exist. These observations would have been strengthened by approaches facilitating the tracking of ILC populations for longer periods of time, but concerns regarding the potential loss of the photoconverted red form of the fluorochromes inhibited such studies. In the future, mice enabling fate mapping of Cre recombinase activity induced through photoactivation might provide a superior approach to the long-term tracking of tissue residency.

Although the IL-7Rα+T-bet+EOMES ILC1 population detected within LNs is developmentally distinct from cNK cells (44), it is notable that the data provided here reveal a similar migratory behavior. Direct comparison of the “helper” ILC1 (17) and cNK cell migration indicated that cNK cells were more migratory because a higher proportion of the cNK cells had newly entered the tissue within 72 hours of labeling. Our data also provide direct evidence that both cNK cells and ILC1s recirculate through peripheral LNs, a behavior demonstrable in photoconvertible mice. Although only a few labeled ILCs were detected within contralateral LNs, this likely reflects the technical limitations of trying to track a population of a few hundred cells within the entire mouse, and the larger cNK cell population was more readily detected, as were T cells. It remains possible, however, that some ILCs exit peripheral LNs and traffic to tissues that were not assessed. The data presented here do not contradict published studies of parabiotic mouse models because here ILCs in the mLN and spleen, but not peripheral LNs, were assessed, and lymphatic migration would not be captured (9). Consistent with these previous studies, our data also indicate that the ILC2 and ILC3 compartments of peripheral LNs contain resident populations. Whereas both the Kaede and H2B-Dendra2 mouse models indicated that the majority of ILC2s and ILC3s were tissue resident, identification of ILC2s by GATA-3 expression in the H2B-Dendra2 mice did identify a sizeable migratory ILC2 population in the LNs (about 40% of total LN ILC2). Few ILC2s are present in LNs compared with tissues such as the lung, and the role of these cells is unclear. Our data indicate that there is some movement of these cells between the circulation and peripheral LNs. The ILC3 subset displayed the least evidence of migratory behavior within the time frame assessed, and it remains possible that the cells present here are the remnants of the lymphoid tissue–inducer population that initially seeded the LN anlagen in the embryo.

The function of the helper ILC populations in LNs is poorly understood, whereas cNK cells have been considered to be key early producers of effector cytokines within the LN to support the initiation of T cell responses. NK cells were initially described as being specifically recruited into draining LNs (42) in response to immunization. Subsequent studies identified that cNK cells enter peripheral LNs via the HEVs in a CD62L- and CCR7-dependent manner (27) and egress dependent on S1P5 (32, 33). Thus, increases in cNK cell numbers in the draining LN may reflect the restricted exit of cells to drive their accumulation, rather than enhanced recruitment. Here, we demonstrate that both resident and migratory ILC1s rapidly produced IFN-γ in response to immunization, as described for cNK cells. Questions remain as to how migratory and resident ILC1s differ in their functions, and whether LN ILC1s contribute to peripheral LNs beyond the roles described for cNK cells. The degree to which ILC1s can contribute to TH1 priming in the absence of cNK cells need to be tested. Given the location of cNK cells within LNs (45), it is likely that ILC1s may reside away from the interfollicular zone and border between the B and T cell areas, where ILC2s and ILC3s have been described (18).

In summary, the data presented here shed new light on the relationship between ILC populations within the circulation and peripheral lymphoid tissues, revealing that a murine ILC1 subset continuously traffics between the blood and peripheral LNs. Despite rapid advances in our understanding of how ILCs develop, much of their basic biology remains poorly understood, and the dynamic changes in LN ILCs described here provide further insight into how these cells might contribute to the regulation of adaptive immunity in lymphoid tissues.

MATERIALS AND METHODS

Study design

The main aims of this study were to understand the extent of ILC migration into and out of peripheral LNs and determine whether ILCs trafficked from skin into LNs. We sought to address these aims through specifically labeling ILCs in peripheral LNs and skin, testing the extent to which ILCs entered and left these tissues and identifying the sites to which these cells migrated. These were done using transgenic mice expressing green photoconvertible proteins, which, when exposed to violet light, switched to a red fluorescence that could then be detected for a number of days after photoconversion. A key feature of the photoconversion performed was that this was optimized such that all cells in the target tissue (e.g., LN or ear skin) simultaneously converted to the red fluorescent form. This enabled both the tracking of labeled red cells migrating to different sites and detection of new cells (expressing the native green form of the photoconvertible) entering the tissue after labeling. The migration of ILCs was assessed under steady-state conditions and in response to inflammation.

Mice

Animal husbandry and experiments were ethically reviewed and carried out in accordance with UK Home Office guidelines. C57BL/6, Kaede (28), H2B-Dendra2 (35), CCR6−/− Kaede, and CCR7−/− Kaede mice were maintained and bred at the University of Birmingham Biomedical Services Unit. Mice were culled between the ages of 6 and 15 weeks.

Experimental procedures

To establish inflammation in the ear (MC903 AD model), 4 nmol of calcipotriol (Tocris Bioscience), in absolute ethanol (VWR Chemicals), was applied to the dorsal and ventral area of the ear for five consecutive days. As a vehicle control, absolute ethanol was used.

Mice were injected intraperitoneally with FTY720 (1 mg/ml in 200 μl of H20; Sigma). As a vehicle control, mice were injected intraperitoneally with H20. When FTY720 was administered in conjunction with bLN surgery or the MC903 model, mice were injected 1 day before the start of the procedure and every day during the procedure.

For photoconversion of the bLN, mice were subcutaneously injected with 100 μl of analgesic buprenorphine 1 hour before surgery (Vetergesic, Reckitt Benckiser Healthcare). Under anesthesia via gaseous isoflurane, a small incision was made exposing the bLN to violet light [365 nm of ultraviolet (UV) light-emitting diode] for 3 min and 30 s. The skin was sutured together, and mice were subcutaneously injected with 500 μl of saline solution. Time points (0 hours) were culled immediately after surgery.

For violet light conversion of the ear, under anesthesia ears were exposed to a violet light (405 nm of UV laser) for 40 s with periodic breaks. Black cardboard was used to shield the rest of the mouse and 0-hour time points were culled immediately after photoconversion.

Immunizations were performed under anesthesia, via gaseous isoflurane, using 2W1S peptide conjugated to OVA protein. The 2W1S peptide is a variant of peptide 52 to 68 from the I-Eα chain that is highly immunogenic in C57BL/6 mice (46). The antigen and adjuvant were administered through intramuscular injections according to GlaxoSmithKline (GSK) standards or through paw pad injections. Upon intramuscular injections, the site of the injection, the gastrocnemius muscle, was shaved, and about 3 mm of the needle was inserted into the muscle. The injection volume of 10 μl was administered into the gastrocnemius muscle of both hind legs. The injections contained 20 μg of OVA-2W1S (ProImmune) and 1:50 human dose AS01B [i.e., 1 μg of each monophosphoryl lipid A (MPL) and QS-21 (Quillaja saponaria Molina, fraction 21)] or 20 μg of OVA-2W1S and PBS. AS01 is a proprietary GSK adjuvant system containing MPL, QS-21, and liposome. QS-21 is licensed by GSK from Antigenics LLC, a wholly owned subsidiary of Agenus Inc., a Delaware, U.S. corporation. For analysis at day 7, a second injection of OVA-2W1S with AS01 or PBS was administered 6 hours before analysis of the response.

Cell culture

In AS01 experiments, assessing the production of IFN-γ, cell suspensions were incubated for 3 hours in culture medium in the presence of brefeldin A (10 μg/ml; BD Biosciences) and monensin (10 μg/ml; BD Biosciences) at 37°C, 5% CO2.

Flow cytometry

Tissues were prepared using the same methods in Dutton et al. (47). LNs were cleaned and teased in RPMI 1640 medium (Thermo Fisher Scientific) and crushed through a 70-μm filter. Samples were centrifuged (5 min, 1500 rpm, 4°C), the supernatant was removed, and samples were resuspended in appropriate amount of Dulbecco’s PBS (DPBS) supplemented with 2% fetal bovine serum (FBS) and 0.5% EDTA. Ears were cut into small sections and agitated while incubated at 37°C in a thermomixer (Eppendorf) for 30 min in 1 ml of Liberase TM/deoxyribonuclease (DNase) digestion solution [3 ml per ear; 3 ml of Dulbecco’s modified Eagle’s medium (DMEM; Thermo Fisher Scientific), 75 μl of Liberase TM (0.28 Wünsch units/ml), and 75 μl of DNase (200 mg/mL)]. Tissue was then filtered through a 70-μm filter and washed with DMEM. The ear was then removed from the filter, and this process was repeated twice more. After the third incubation, the ear was crushed through the filter and washed with DMEM. Samples were centrifuged and suspended in an appropriate amount of DPBS supplemented with 2% FBS and 0.5% EDTA.Blood (1 to 2 ml) was harvested via cardiac puncture. Samples were centrifuged (5 min, 2000 rpm), the supernatant was removed, and samples were incubated in 500 μl of Gey’s solution (47) on ice for 5 min. This was repeated twice more, before being resuspended in appropriate amount of DPBS supplemented with 2% FBS and 0.5% EDTA. Samples enriched for T and B cells were stained for B220 [allophycocyanin (APC)], CD3 (APC), and CD5 (APC) for 30 min on ice, followed by an enrichment with anti-APC MicroBeads [magnetic-activated cell sorting (MACS)]. Samples enriched for CD45+ hematopoietic cells were enriched using CD45 MicroBeads (MACS). All samples were stained with a LIVE/DEAD cell viability assay in APC-Cy7 in PBS (1:1000) for 15 min at 4°C. Extracellular staining was conducted with antibodies diluted in DPBS supplemented with 2% FBS and 0.5% EDTA for 30 min on ice. The Foxp3 fixation and permeabilization kit (eBioscience) was used to fix and intracellularly stain samples. Kaede samples did not undergo fixation and intracellular staining. The antibodies raised against the following mouse agents were used: B220 (1:300 or 1:200; clone RA3-6B2, eBioscience), CCR6 (1:100; clone 29-2 L17, BioLegend), CCR7 (1:100; clone 4B12, eBioscience), CD3ε (1:100 or 1:200; clone 145-2C11 or eBio500A2 or 17A2; eBioscience, BD Biosciences, or BioLegend), CD4 (1:100; clone RM4-5, BioLegend), CD5 (1:100 or 1:200; clone 53-7.3, eBioscience), CD11b (1:300 or 1:200; clone M1/70, eBioscience), CD11c (1:300 or 1:200; clone N418, eBioscience or BioLegend), CD19 (1:200; clone eBio1D3, eBioscience), CD25 (1:50; clone PC61, eBioscience), CD31 (1:400; clone 390, eBioscience), CD45.2 (1:100; clone 104, BioLegend), CD49b (1:200; clone DX5, eBioscience), CD62L (1:1500, 1:500, or 1:100; clone MEL/14, eBioscience or BioLegend), CD69 (1:100; clone H1.2F3, eBioscience), CD103 (1:200; clone 2E7, BioLegend) CD123 (1:500; clone 5B11, eBioscience), CD207 (1:100; clone 4C7, BioLegend), c-kit (1:100; clone H57-597, eBioscience), EOMES (1:50; clone Dan11mag, eBioscience), F4/80 (1:200; clone BM8, eBioscience), FcεRI (1:200; clone MAR-1, eBioscience), GATA-3 (1:75; clone TWAJ, eBioscience), gp38 (1:1000; clone 8.1.1, BioLegend), Gr-1 (1:2000; clone RB6-8C5, eBioscience), IL-7Rα (1:50; clone A7R34, BioLegend), Ki-67 (1:400; clone SolA15, eBioscience), KLRG1 (1:200; clone 2F1, BioLegend or eBioscience), MHCII (1:100; clone M5/114.15.2, eBioscience), NK1.1 (1:100 or 1:200; clone PK136, eBioscience or BD Biosciences) NKp46 (1:50 or 1:100; clone 29A1.4, BioLegend or eBioscience), RORγ (1:100; clone AFKJS-9, eBioscience), T-bet (1:50; clone eBio4B10, eBioscience), T cell receptor β (TCRβ; 1:100; clone H57-597, BioLegend), TCRγδ (1:200; clone GL3, BD Biosciences), and TER119 (1:200 or 1:100; clone TER-119, eBioscience). To identify ILCs, the lineage channel included B220, CD11b, CD11c, CD3, CD5, CD19, TER119, Gr-1, CD123, F4/80, FcεRI, and CD49b. CD123 was not included in the lineage channel when identifying ILCs in Kaede mice. Samples were run on the BD LSRFortessa X-20 (BD Biosciences), and data were collected using BD FACSDiva software (BD Biosciences).

Gene expression analysis

FACS-sorted NK1.1+/− ILCs were lysed in RLT buffer (Qiagen), and total RNA was subsequently isolated using a Quick-RNA MicroPrep kit according to the manufacturer’s instructions (Zymo Research). Complementary DNA was generated from total RNA in a reverse transcription reaction using random primers. TaqMan gene expression assays for selected genes [S1pr1 (Mm00514644_m1), S1pr2 (custom plus TaqMan RNA assay) S1pr3 (Mm04229896_m1), S1pr4 (Mm00468695_s1), and S1pr5 (Mm02620565_s1); Applied Biosystems] were prepared using the Taqman PreAmp Master Mix Kit. Samples were then analyzed on QuantStudio 12 K Flex RT-PCR system. Quantitative polymerase chain reaction results were analyzed using the R environment for statistical computing. ΔΔCt values were calculated using the mean of three reference genes (β-actin, glyceraldehyde 3-phosphate dehydrogenase, and 18S ribosomal DNA). Fold changes were determined by calculating power 2-ΔΔCt.

Statistical analysis

Data collected were analyzed using FlowJo 10.4.2 software (Treestar) and GraphPad Prism 7. Normal distribution of the data was not assumed. Pairs of samples were compared using an unpaired two-tailed Mann-Whitney U test. When comparing more than two sets of data, statistical significance was tested using Kruskal-Wallis one-way analysis of variance (ANOVA) with post hoc Dunn’s test: *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001. The bar represents the median, and where appropriate, the median has been shown numerically. Where a P value is not indicated, no statistical difference was observed.

SUPPLEMENTARY MATERIALS

immunology.sciencemag.org/cgi/content/full/4/35/eaau8082/DC1

Fig. S1. LN stromal populations remain Kaede Red+ after photoconversion.

Fig. S2. ILCs continuously traffic into LNs with different kinetics to T cells.

Fig. S3. Photoconversion does not cause an increase in the number of ILCs within the labeled bLN.

Fig. S4. Expression of S1P1 by ILC subsets in the bLN.

Fig. S5. Treatment with FTY720 results in ILC accumulation within the bLN.

Fig. S6. Analysis of S1PR expression by ILCs.

Fig. S7. Identification of ILC subsets within the bLN.

Fig. S8. Analysis of ILC residency in LNs using H2B-Dendra2 mice.

Fig. S9. Differences in Dendra Red expression by ILC2s dependent on identification by a TF or cell surface marker expression.

Fig. S10. Expression of Ki-67 by ILC subsets within the bLN.

Fig. S11. Comparison of ILC1 and NK cell recirculation through peripheral lymph nodes.

Fig. S12. Induction of atopic dermatitis causes an increase in the frequency of ILC2s in ear skin.

Fig. S13. CCR6 is expressed by all ILC subsets in the ear and auLN.

Table S1. Raw data.

REFERENCES AND NOTES

Acknowledgments: The 2W1S:I-Ab tetramer was obtained from the NIH Tetramer Facility. We thank Y. Miwa (Tsukuba University) and O. Kanagawa (RCAI, RIKEN) for the Kaede mice. We thank O. Pabst (RWTH Aachen University) for providing the H2B-Dendra2 mice. We thank the Biomedical Services Unit at the University of Birmingham for all the help with in vivo experimental work. We are very grateful to M. Hepworth and S. Milling for critical reading of the manuscript. Funding: This work was supported by a Senior Research Fellowship from the Wellcome Trust to D.R.W. (grant number 110199/Z/15/Z). E.E.D. is a BBSRC CASE student sponsored by MedImmune LLC, who partially funded the studentship (BB/L016893/1).Author contributions: E.E.D. designed and performed experiments, analyzed data, performed statistical analysis, and wrote the manuscript; D.W.G., C.W., R.F., and E.L.B. designed and performed experiments; F.P. and C.A.M. performed experiments and analyzed data; A.C., M.A.S., and G.C. designed experiments and analyzed and critiqued data; M.C. and A.M.D. provided key reagents, designed experiments, and critiqued data; M.T. provided key reagents and expert advice in experimental design and methodology; D.R.W. designed and performed experiments, analyzed data, performed statistical analysis, and wrote the manuscript. Competing interests: A.C., G.C., F.P., and C.A.M. are full-time employees of MedImmune LLC. M.A.S. is a former employee of MedImmune and a full-time employee of Regeneron. All past or current MedImmune coauthors own or owned Astra Zeneca stock and/or stock options. M.C. and A.M.D. are full-time employees of the GSK group of companies. A.M.D. reports ownership of shares and/or restricted shares in GSK. The other authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials.
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