Vitamin A Controls the Presence of RORγ⁺ Innate Lymphoid Cells and Lymphoid Tissue in the Small Intestine

The intestinal immune system is an extensive network of immune cells and lymphoid tissues, including Peyer’s patches, cryptopatches (CPs), and isolated lymphoid follicles (ILFs) (1). Although the immune cells, which are present within these lymphoid tissues or in the lamina propria, are protected by a dense mucus layer, foreign particles such as commensal bacteria or diet-derived products are in constant close proximity. To prevent unnecessary inflammatory reactions, most gut immune cells differentiate to become unresponsive toward compounds commonly found in the gut. However, these cells still need to mount an efficient immune response against invading pathogens. Therefore, the gut immune system is tightly regulated (2).

The delicate balance in the intestines is maintained by various immune cells of which innate lymphoid cells (ILCs) have recently been shown to play a crucial role in lymphoid tissue organogenesis, maintenance of epithelial integrity, and tissue remodeling and repair. ILCs originate from a common innate lymphoid progenitor that emerges from the common lymphoid progenitor both in the bone marrow and fetal liver (1, 3). Furthermore, ILCs have a lymphoid morphology, lack myeloid and dendritic cell markers, and do not possess recombinant activating gene–dependent rearranged Ag receptors, which distinguishes them from T and B lymphocytes (4). During their development, ILCs depend on common cytokine receptor–γ-chain signaling and the expression of DNA inhibitor Id2 (3–5). Finally, ILCs are essential components in early stages of innate immunity against microbes (1, 3).

Based on distinct cytokine profiles and transcriptional requirements, ILCs have been classified into group 1, group 2, and group 3 ILCs. The different ILC types seem to mirror the Th cell subsets and are classified according to their function and phenotype due to the variability in cytokine profile (6). NK cells are the prototypical group 1 ILCs and are characterized by the production of IFN-γ and their cytotoxic functions. Besides the NK cells, ILC1 cells are also members of the group 1 ILCs. However, ILC1 cells are able to originate from group 3 ILCs, and it is therefore hypothesized that they may represent a distinct cell type (6, 7). The second group is formed by type 2 ILCs, which are controlled by the transcription factors retinoic acid–related orphan receptor (ROR)α and GATA3 for their differentiation. They are able to produce type 2 cytokines such as IL-5 and IL-13 and therefore play an important role during helminth infection and allergic lung inflammation (8). Additionally, within the group 3 ILCs there are three different subsets described that have the capacity to produce IL-17 and IL-22 cytokines. First, this group includes the lymphoid tissue inducer (LTi) cells, which have a crucial role in embryonic lymph node formation (9). Furthermore, two other subsets fall within group 3 ILCs, which both express the transcription factor RORγ, similar to LTi cells. These two subsets can be discriminated by the expression of NCR receptors. Whereas NCR⁺ ILC3s only produce high amounts of IL-22, NCR⁻ ILC3s produce in addition IL-17 and IFN-γ (7, 10). Group 3 ILCs are now being recognized for their essential role in both the formation of gut-associated lymphoid tissue and in intestinal host defense (5, 6).

Recent findings have suggested that commensal bacteria and nutritional components can influence ILC populations within the gut. However, the identification of these particles is impeded by the enormous diversity of nutrients and by indirect sources such as bacteria capable of transforming compounds into a form that can be digested by the host. For instance, it has been shown that dietary AhR ligands are able to control the function and maintenance of group 3 ILCs postnatally, which consequently has an impact on the formation of CPs and isolated lymphoid follicles (11–13). Additionally, it was first shown by Mielke et al. (14) that not only intestinal γδ T cells but also IL-22–producing ILCs are controlled by the active metabolite of vitamin A, retinoic acid (RA). The importance of vitamin A for ILCs was even further established, as in the absence of vitamin A RORγ⁺ ILC3s are reduced, whereas group 2 ILCs are enhanced (15). Moreover, fetal LTi differentiation and immune fitness of adult mice were shown to be determined by the available levels of maternal retinoids during pregnancy (16). These levels of maternal retinoids affect the number of fetal ILC3s and therefore control the size of secondary lymphoid tissue, which are formed prenatally. Consequently, the levels of prenatal vitamin A affect the efficiency of immune responses mounted in these animals during adulthood (16). However, the direct role of RA on the development of the different ILC3 subsets and lymphoid tissue postnatally and in adult life has not been well studied.

In this study, we examine the role of the RA on the ILC3 subsets within the small intestine postnatally and in adult life. We show that vitamin A deficiency results in a specific reduction of RORγ⁺ ILC3s, whereas paradoxically only the NCR⁻RORγ⁺ ILC3 subset was enhanced upon addition of RA in vivo. Furthermore, we were able to show that also the numbers of CPs and ILFs, collectively called solitary intestinal lymphoid tissues (SILTs), were reduced in vitamin A–deficient (VAD) mice. Moreover, upon postnatal blockade of RA signaling, using the pan-RA receptor antagonist BMS 493 (17), defects in SILT formation were observed. To demonstrate that RA signaling has a direct effect on these RORγ⁺ ILC3s, we crossed RORγ-cre mice with RARα-DN mice, leading to impaired RA signaling in RORγ⁺ cells. These mice showed a specific reduction of NCR⁻ and NCR⁺ ILC3, but not LTi cells, within the small intestine. Consequently, the formation of enteric lymphoid tissue was not affected in these mice. In conclusion, our data further define the role of vitamin A in the regulation of intestinal ILC3s and the formation of intestinal lymphoid tissue postnatally.

C57BL/6 mice, obtained from Charles River Laboratories (Maastricht, the Netherlands), were mated overnight, and the morning of vaginal plug detection was marked as embryonic day (E)0.5. Starting at E8–E9, pregnant females either received a chemically defined diet that lacked vitamin A (the modified AIN-93M feed; MP Biomedicals, Santa Ana, CA; VAD) or contained vitamin A (4000 IU/kg in the modified AIN-93M feed; MP Biomedicals; vitamin A control [VAC]) with use of vitamin-free casein (18). All mice were housed under specific pathogen-free conditions. Pups were weaned at 4 wk of age and maintained on the same diet until at least 10 wk of age, when the analyses were performed. For generation of RA^high mice, C57BL/6 adult mice (12–14 wk of age) were fed with global 16% protein rodent diet (Harlan, Horst, the Netherlands), which contains 4.5 μg RA per gram dry food supplemented with 100 μg RA per gram dry food or vehicle control for 1–2 wk (Sigma-Aldrich, Zwijndrecht, the Netherlands). Diet was refreshed twice a day to reduce the effects of RA degradation by light. For in vivo blockade of RA signaling, mice were treated with the pan-RAR antagonist BMS493 (5 mg/kg; Tocris Bioscience) or vehicle (DMSO) in corn oil (16, 19). Mice were treated for 7 d via oral gavage twice a day with 10- to 12-h intervals starting at day 4 after birth. After 7 d, mice were sacrificed for analysis or left untreated for another 21 d, after which they were sacrificed for analysis. RORγt-cre mice (20) were crossed with ROSA26-RARα403 (21) mice to abrogate RA signaling in specific RORγ⁺ cells.

All animal experiments were approved by national and institutional ethical committees, including the Direção Geral de Veterinária and Instituto de Medicina Molecular Ethical Committee and the Animal Experimental Committee, VU University Medical Center.

Small intestines were dissected and opened longitudinally after removal of Peyer’s patches. Small intestines were washed with HBSS without Ca²⁺ and Mg²⁺ (Invitrogen, Carlsbad, CA) containing 15 mM HEPES (Invitrogen) and 250 μg/ml gentamicin to remove fecal contents. Small intestinal segments were incubated twice with HBSS (Invitrogen) containing 5 mM EDTA (Sigma-Aldrich), 15 mM HEPES (Invitrogen), 10% FCS (HyClone Laboratories/Greiner Bio-One, Alphen aan den Rijn, the Netherlands), 1 μM DTT (Promega Benelux, Leiden, the Netherlands), and 14 mM 2-ME (Sigma-Aldrich) for 15 min at 37°C while constantly stirring to remove mucus. Pieces of small intestine were further cut with scissors and digested at 37°C for 20 min, using 150 μg/ml Liberase Blendzyme 2 (Roche, Penzberg, Germany) and 200 μg/ml DNAse I (Roche) in HBSS (Invitrogen) containing 15 mM HEPES (Invitrogen) and 10% FCS (HyClone Laboratories/Greiner Bio-One) while constantly stirring. The cell suspensions were filtered through 70-μm cell strainers (BD Biosciences, Breda, the Netherlands) and the recovered cells were washed twice with HBSS without Ca²⁺ and Mg²⁺ (Invitrogen) containing 15 mM HEPES (Invitrogen). CD45⁺ cells were positively selected using PE-Cy7–labeled anti-CD45 (clone 30-F11; eBioscience/Immunosource, Halle-Zoersel, Belgium) and the EasySep PE positive selection kit (StemCell Technologies, Grenoble, France). Purified CD45⁺ lamina propria cells were used for either RNA isolation, flow cytometry and analyzed with a CyAn ADP flow cytometer (Beckman Coulter) or for ex vivo culture with a GolgiPlug (brefeldin A, BD Biosciences) for 4 h.

Tissues were embedded and frozen in OCT compound (Sakura Finetek Europe, Zoeterwoude, the Netherlands). Total small intestine tissue was sectioned by 7-μm-thick cryosections, which were all collected. Every 10th slide was stained for screening assay to count and characterize all SILTs present within the full small intestine. Sections were fixed in acetone for 5 min and air dried for an additional 10 min, followed by preincubation with PBS supplemented with 10% (v/v) mouse serum for 15 min. Incubation with directly conjugated Abs was performed for 30 min. All incubations were performed at room temperature. Stainings were analyzed on a Leica DM6000 microscope (Leica Microsystems Nederland, Rijswijk, the Netherlands).

Anti–CD4-555 (clone GK1.5), anti–B220-488 (clone 6b2), and anti–CD45-647 (clone MP33) were used for immunofluorescence, which were all affinity purified from hybridoma cell culture supernatants with protein G–Sepharose (Pharmacia, Uppsala, Sweden) and coupled to Alexa Fluor labels (Invitrogen).

For flow cytometry analysis, anti–CD45-PE-Cy7, anti–CD4-PerCP-Cy5.5, anti–CD3-488, ant–Ki-67-647, anti–RORγ-PE, and anti–NKp-46 (all eBioscience, San Diego, CA) were used. Intracellular staining for RORγ was performed with the FoxP3 staining kit (eBioscience). Live/Dead fixable near-IR stain fluorescence (Invitrogen) was used to exclude dead cells.

RNA was extracted from total homogenized small intestine or from CD45 selected cells using TRIzol reagent (Invitrogen). RNA was isolated by precipitation with isopropanol. cDNA was synthesized from total RNA using a RevertAid first-strand cDNA synthesis kit (Fermentas Life Science, St. Leon-Rot, Germany) according to the manufacturer’s protocol. Quantitative real-time PCR was performed on an ABI Prism 7900 sequence detection system (Applied Biosystems, Foster City, CA). The reaction mixture was composed of SYBR Green master mix (Applied Biosystems), 300 nM each primer, and cDNA with a total volume of 10 μl, according to the manufacturer’s instructions. Primers were designed across exon–intron boundaries using Primer Express software and guidelines (Applied Biosystems); the primer sequences can be found in Table I. For IL-17 a primer set (Mm00439619, Life Technologies, Carlsbad, CA) was ordered and detected with TaqMan (Applied Biosystems).

Using the comparative C_T method (ΔC_T), relative changes in mRNA levels between samples were determined.

Feces from the small intestines of mice were collected in cold PBS buffer. Debris was removed by cold centrifugation for 20 min at 2000 rpm to harvest the supernatant for analysis of secretory IgA. Supernatants of mesenteric lymph node dendritic cells and B cell cocultures were used directly for ELISA. Plates were coated with anti-mouse IgA Ab to capture secretory IgA or mouse IgA used as a standard (clone s107), followed by anti-mouse IgA-biotin Ab (all SouthernBiotech, Birmingham, AL) and subsequently streptavidin labeled with HRP. Samples were analyzed with a FLUOstar Optima microplate reader (BMG Labtech, Isogen Lifescience, de Meern, the Netherlands).

Results are given as the mean ± SEM. Statistical analyses were performed using GraphPad Prism 4 (GraphPad Software, La Jolla, CA) with either the two-tailed Student t test or two-Way ANOVA with a Bonferonni correction. A p value <0.05 was considered significant.

To establish the role of dietary-derived products on the regulation of intestinal ILCs we made use of a VAD mouse model described by Iwata et al. (18). We analyzed the small intestines by flow cytometry for the expression of RORγ and the NCR receptor NKp46, gated on live CD45⁺CD3⁻ cells. The percentages and cell numbers of both NKp46⁺RORγ⁺ and NKp46⁻RORγ⁺ cells were significantly reduced in the absences of vitamin A, whereas NKp46⁺RORγ⁻ cells from the ILC1 group were equally represented in both groups (Fig. 1A, Supplemental Fig. 1A). This effect was only seen when vitamin A was completely absent, as mice receiving a vitamin A-low diet, resulting in vitamin A plasma levels just above the detection limit (data not shown), did not show this reduction (Fig. 1A). Only the cell numbers of double-positive NKp46⁺RORγ⁺ ILC3s were slightly reduced in mice with a vitamin A-low diet (Supplemental Fig. 1A). Based on Ki-67 expression, we observed an increased proliferation in both RORγ⁻ as well as RORγ⁺ intestinal cells, gated on CD45⁺CD3⁻ in VAD animals compared with control mice (Supplemental Fig. 1B). These data suggested that vitamin A can inhibit proliferation and thereby might induce specific differentiation in all CD45⁺CD3⁻ intestinal cells. Whereas significant effects were shown within the intestines, no differences for NKp46⁺NK1.1⁻ ILC3s and NKp46⁺NK1.1⁺ ILC1s were found in the mesenteric and peripheral lymph nodes of VAC versus VAD mice (Supplemental Fig. 1C), suggesting the effect of vitamin A metabolites on ILC differentiation takes place within the intestine.

To address whether we could specifically stimulate group 3 ILC differentiation within the small intestines, we supplemented conventional rodent diet with RA or a vehicle control for 1–2 wk. In contrast to the VAD mice in which NCR⁻ and NCR⁺ ILC3s were affected, mice that received additional RA showed a specific increase of only NKp46⁻RORγ⁺ ILC cells (Fig. 1B, Supplemental Fig. 1D). Additionally, in both in vivo models group 1 ILCs, which are NKp46⁺ but RORγ⁻, showed no differences compared with control mice (Fig. 1A, 1B).

Group 3 ILCs are important producers of IL-17 and IL-22 cytokines in the lamina propria of the gut (1, 5), and we therefore explored whether these cytokines were altered upon vitamin A deficiency. We found diminished percentages of CD3⁻IL-22⁺ cells in VAD mice when compared with control mice (Fig. 1C, Supplemental Fig. 1E). Furthermore, a lower mRNA expression of IL-22 and IL-17 was detected in CD45⁺ cells isolated from the lamina propria of VAD mice (Fig. 1D). The production of IL-22 is crucial for inducing the secretion of antimicrobial defensins by epithelial cells at the luminal side of the intestines (1, 22). Therefore, we investigated the mRNA levels of the defensins RegIIIβ and RegIIIγ within the total small intestine along the gut axis divided in three equal parts from proximal to distal side. The middle part of the small intestine of VAD mice displayed a significant decrease in mRNA levels of both RegIIIβ and RegIIIγ compared with VAC intestines (Fig. 1E). These data correlate with the differences we obtained for the expression of IL-22 in VAD mice. Alternatively, we obtained significant higher levels of RegIIIγ within the distal part of the small intestine after supplementation with RA, whereas we demonstrated only a trend of induced IL-17, IL-22, and RegIIIβ mRNA levels in the small intestine upon RA stimulation (Supplemental Fig. 2).

In conclusion, our data further substantiated the role of vitamin A in regulating intestinal group 3 ILCs. Additionally, we showed that the related cytokines IL-17 and IL-22 were affected as well, which correlated with changed expression of defensins by the intestinal epithelial cells.

Because group 3 ILCs also include LTi cells, we investigated whether the development of intestinal lymphoid tissue was affected in mice raised on a VAD diet. In this model vitamin A is withheld from the diet given to pregnant mice starting at E8.5–E10.5. Therefore, the offspring will only become completely deficient by the adult age of 10 wk (23). Although Peyer’s patches develop already before birth, SILTs are generated postnatally (24, 25). To quantify these lymphoid tissues, total small intestine was screened for SILTs and they were characterized by both size and phenotype. We discriminated between CPs, which mainly contain RORγ⁺CD4⁺ LTi cells, immature ILFs (iILFs) harboring some B cells, and mature ILFs (mILFs), which consist of a fully developed B cell follicle. Whereas we could detect all different stages of SILT clusters in both VAC and VAD mice (Fig. 2A), we did observe reduced numbers of SILTs in both the middle and distal part of VAD small intestines (Fig. 2B). Upon further analysis, the different stages of these SILTs, that is, CP, iILF, and mILF, were similarly represented in the control and deficient group (Fig. 2B). When we divided the SILTs according to their size in five different categories described by Pabst et al. (26), again no differences were obtained within the distribution between the different mice (Fig. 2C), indicating that SILT numbers, and not their maturation, are affected. Although there is not much difference in SILT size, the average size in VAD mice was significant larger compared with the SILTs present in VAC mice (Fig. 2C). Nevertheless, the IgA concentrations measured in the small intestine luminal content are significantly lower in VAD mice compared with control mice (Fig. 2D), demonstrating the functional effects of the reduction in SILTs. In contrast to the effects on SILT formation, no differences could be observed in the prenatally developed Peyer’s patches, as their numbers were similar in both groups (Fig. 2E).

Taken together, these data show that upon vitamin A deficiency fewer SILTs develop probably as the result of reduced numbers of RORγ⁺ ILCs within the gut. Furthermore, the SILTs that develop showed no abnormalities with respect to size and phenotype when compared with SILTs in control intestines. Thus, the observed reduction in total IgA levels within the small intestinal lumen are likely caused by the diminished number of SILTs present in VAD mice.

Although our VAD model already indicated an important role for the induction of group 3 ILCs and consequently the formation of SILTs by vitamin A postnatally, this model does not allow immediate abrogation of available RA levels. Therefore, we used an in vivo model where we blocked signaling of RA receptors shortly after birth with the use of the pan-RA receptor antagonist BMS 493 (17). Mice were treated twice a day by oral gavage with either DMSO control or BMS 493 for 7 d. The experiment was started 4 d after birth and maintained until day 11 of age. Flow cytometry analysis showed that most RORγ⁺ ILCs were still NCR⁻ in the small intestine of these young mice, whereas both percentages of NKp46⁻RORγ⁺ and NKp46⁺RORγ⁺ cells were significantly reduced after treatment with BMS 493 (Fig. 3A). Additionally, we demonstrated that within the RORγ⁺ group 3 ILCs both CD4⁻ and CD4⁺ are equally represented in DMSO-treated mice, indicating that the small intestines at this age still harbor a significant population of CD4⁺RORγ⁺ LTi cells. However, BMS 493 treatment resulted in significant reduction in the percentages of CD4⁻ as well as CD4⁺RORγ⁺ ILCs. Accordingly, when small intestines were analyzed 3 wk after the treatment was stopped, remarkable differences in SILT development between the DMSO- and BMS 493–treated mice were noticed. Within the DMSO-treated mice, SILTs consisted of LTi CD4⁺ cells and some B cells could be found, although B cell follicles were absent (Fig. 3B). However, in mice treated with BMS 493 we could only observe a few small clusters of LTi CD4⁺ cells, which never contained B cells (Fig. 3C). Additionally, the clusters found in BMS 493–treated mice were much smaller compared with the SILTs that developed in control intestines (Fig. 3C). Similar as seen in VAD mice, BMS 493–treated mice did not show any differences in Peyer’s patch formation compared with control mice (Fig. 3C).

Collectively, these data suggest that vitamin A–mediated signaling affects the group 3 ILCs in the small intestine postnatally, which consequently influences SILT formation within the lamina propria.

To demonstrate that RA has a direct effect on the differentiation of group 3 ILCs, we made use of a more specific model. We crossed RORγ-cre mice with RARαDN mice, resulting in an overexpression of human RARα, which lacks an intracellular signaling domain, in RORγ⁺ cells. The overexpression of this nonsignaling receptor will prevent RA signaling by the normal expressed RA receptors on RORγ⁺ cells (27). Analysis of the small intestine by flow cytometry showed a significant reduction of NKp46⁻RORγ⁺ as well as NKp46⁺RORγ⁺ group 3 ILCs in RORγ-RARαDN mice (Fig. 4A–C). Additionally, analyzing the mean fluorescence intensity of RORγ, we detected a significantly lower signal in all RORγ⁺ group 3 ILCs in RORγ-RARαDN mice compared with control mice, whereas NKp46⁺RORγ⁻ ILCs were not affected (Fig. 4D). Remarkably, however, LTi cells, which are necessary for lymphoid tissue formation, were not changed in RORγ-RARαDN mice (Supplemental Fig. 3C). Subsequent analysis of the SILTs within these mice also revealed that the SILT number as well as SILT phenotype were not different from control mice (Supplemental Fig. 3A, 3B). Additionally, the levels of IgA remained similar within the two different mice (Supplemental Fig. 3D). Our data further confirm our earlier observations that in embryonic RORγ-cre × RARαDN mice the RORγ expression, which is needed to induce RARαDN expression, is not expressed in time in LTi cells to affect their differentiation (16). We therefore conclude that in RORγ-cre × RARαDN mice, RORγ⁺ group 3 ILCs, excluding LTi cells, are affected. Taken together, our findings demonstrate that RA signaling in RORγ⁺ cells is needed for a proper development of ILCs in the group 3 subfamily.

An increasing number of studies have focused on the effect of dietary components on the immune system and are revealing the mechanisms involved in this interaction. In the present study, we show that dietary vitamin A regulates the postnatal differentiation of intestinal ILCs and SILT formation. When mice were devoid of vitamin A, reduced percentages and numbers of both the NCR⁺ as NCR⁻ group 3 ILCs were present in the small intestine. Moreover, when mice received additional dietary RA the population of NCR⁻ group 3 ILCs was increased specifically when compared with control mice. These data suggest that NCR⁺ ILC3s, which may arise from NCR⁻ ILC3s, need a different trigger than RA for the induction of NCR expression. Although the role of vitamin A and its active metabolite RA on intestinal ILCs has recently been reported (15), no discrimination between NCR⁺ and NCR⁻ group 3 ILCs was made. We additionally demonstrated that RA signaling has a direct effect on RORγ⁺ cells, because genetically abrogation of RA signaling in RORγ⁺ cells resulted in reduced levels of NCR⁻ and NCR⁺ ILC3s.

We showed that vitamin A deficiency resulted in a reduced number of SILTs when compared with mice that were raised on a diet containing normal levels of vitamin A. One has to take into account that in our model, mice have detectable circulating levels of vitamin A and that they will only become fully deficient at adult age. Therefore, a more pronounced reduction in the number of SILTs was observed when we inhibited RA signaling after birth, during the time when SILT formation takes place.

Recently we have demonstrated a crucial role for vitamin A in the prenatal development of secondary lymphoid organs. We have shown that the size of these lymphoid organs determined the efficiency by which a viral infection could be cleared at adult age (16). An essential role for vitamin A in adult mice during Citrobacter rodentium infection and dextran sodium sulfate–induced colitis has been shown (14, 15), indicating an important role for IL-22 production during intestinal inflammation. However, these studies did not consider the reduced number of SILTs in the absence of dietary intake of vitamin A, which will also have its effect on the efficiency of the immune response. Therefore, within these inflammation models not only the reduced production of IL-22 but also the diminished numbers of SILTs and reduced production of IgA may have an effect.

Furthermore, it was shown that RA is able to induce the production of IL-22 binding protein by dendritic cells (28). This binding protein is a soluble receptor for IL-22, which has a higher affinity compared with the membrane-bound receptor and functions as a decoy receptor that controls the activity of IL-22 (29). This control is important, as the production of IL-22 can be beneficial but also harmful to the host. Whereas IL-22 showed protective effects during the first phase of dextran sodium sulfate induced colitis, it can become harmful during the recovery phase (30, 31). It was shown that uncontrolled IL-22 production can lead to the induction of tumor growth of intestinal epithelial cells, which can be suppressed by the production of IL-22 binding protein (31, 32). Moreover, intestinal myeloid cells are responsible for the IL-22 production by ILCs during an active immune response (33, 34). This IL-22 secretion by ILCs is induced upon the production of IL-23 and IL-1β (34, 35). Alternatively, a correlation between the susceptibility of inflammatory bowel disease and the IL-23R gene has been demonstrated, showing that a defect in IL-23 signaling can lead to intestinal inflammation (36, 37). Alternatively, several inflammatory diseases, including inflammatory bowel disease, are shown to have higher levels of IL-22 in humans (22). Therefore, when turned on at the right moment, this signaling pathway provides protection for the host, which makes natural stimulations for this pathway, that is, dietary components, a good strategy for safeguarding mucosal immune homeostasis.

In conclusion, our study demonstrates an essential role for vitamin A in the development of intestinal group 3 ILCs. Additionally, the reduced differentiation of RORγ⁺ ILCs, including LTi cells, led to a decreased development of intestinal lymphoid tissue during vitamin A deficiency or blocking RA signaling. Importantly, we were able to show that RA has a direct effect on intestinal RORγ⁺ ILCs in transgenic mice. With these data, we demonstrated a crucial role for the active metabolite of vitamin A in the development of intestinal group 3 ILCs and postnatally formed lymphoid tissue within the intestine.

The authors have no financial conflicts of interest.