Th17 cell responses orchestrate immunity against extracellular pathogens but also underlie autoimmune disease pathogenesis. In this study, we uncovered a distinct and critical role for miR-18a in limiting Th17 cell differentiation. miR-18a was the most dynamically upregulated microRNA of the miR-17–92 cluster in activated T cells. miR-18a deficiency enhanced CCR6+ RAR-related orphan receptor (ROR)γt+ Th17 cell differentiation in vitro and increased the number of tissue Th17 cells expressing CCR6, RORγt, and IL-17A in airway inflammation models in vivo. Sequence-specific miR-18 inhibitors increased CCR6 and RORγt expression in mouse and human CD4+ T cells, revealing functional conservation. miR-18a directly targeted Smad4, Hif1a, and Rora, all key transcription factors in the Th17 cell gene-expression program. These findings indicate that activating signals influence the outcome of Th cell differentiation via differential regulation of mature microRNAs within a common cluster.

T helper cells orchestrate cellular and Ab-mediated immunity against a variety of pathogens. To accommodate this functional diversity, Th cells differentiate into distinct subsets that are classified by defined phenotypic characteristics and specialized functions in immunity (1). Th17 cells combat extracellular bacteria and fungi, but they have also been implicated in the pathogenesis of several autoimmune and inflammatory diseases, including multiple sclerosis and psoriasis (24). Th17 cells also mediate immune responses that are involved in maintaining epithelial barrier integrity, and it has been widely suggested that some cases of asthma may be caused by dysregulated Th17 responses (4, 5). Studies of human asthma and mouse models indicate that lung inflammation can shift to a response marked by Th17 cytokines and neutrophil infiltration when Th2 cell–mediated eosinophilia is reduced (6, 7). Understanding how Th17 cells are programmed and contribute to tissue inflammation remains an important research frontier with high potential for therapeutic impact in a broad spectrum of inflammatory diseases.

Th17 cells can be generated in vitro by activating CD4+ T cells in the presence of TGF-β and IL-6 (810). They are characterized by IL-17 secretion and expression of their lineage-defining transcription factor, RAR-related orphan receptor (ROR)γt. The closely related factor RORα is also an important transcriptional regulator of the Th17 gene-expression program (11). Th17 cells are commonly identified by expression of the RORγt/RORα target gene Ccr6, which encodes a chemokine receptor important for Th17 cell trafficking into the brain and mucosal tissues (1215). In addition to RORγt and RORα, Th17 cells use multiple other transcription factors in a robust transcriptional network that promotes their gene-expression program (16, 17). Of note, HIF1α is a limiting factor that regulates Th17 cell biology (1820), and SMAD4 cooperates with SMAD2/3 to transduce TGF-β signaling in differentiating Th17 cells.

MicroRNAs (miRNAs) have emerged as potent regulators of Th cell differentiation, function, and plasticity (21). miRNAs are small endogenously expressed RNAs that regulate gene expression at the posttranscriptional level. Individual miRNAs can target hundreds of distinct mRNAs, and each mRNA can have several miRNA binding sites. Th17 cell differentiation is impaired in miRNA-deficient T cells (22), and important roles have been identified for several particular miRNAs (18, 23, 24). Some of these miRNAs indirectly influence Th17 lineage commitment by acting on other cell types (25), whereas others act directly on cell-intrinsic signaling that induces Th17 cell programming, expansion, and effector function in vitro and in mouse models of autoimmunity (18, 2529).

The Mirc1 locus, better known as the miR-17–92 cluster, encodes six miRNAs in four families (miR-17, miR-18, miR-19, and miR-92 families), each defined by a common seed sequence and predicted target genes (30). The miR-17–92 cluster and miRNAs in these four families are important for T cell proliferation and survival, as well as for the proper differentiation and immunological functions of regulatory T, T follicular helper (Tfh), Th1, Th2, and Th17 cells (21, 3141). In Tfh cells, miR-17–92 deficiency also induced inappropriate expression of Th17-associated genes (34). Studies that dissected the functionally relevant miRNAs within the miR-17–92 cluster in T cells have focused almost entirely on the miR-17 and miR-19 families, and they uncovered similar roles in promoting clonal expansion and cytokine production in a variety of Th subsets (31, 32, 35, 40, 41). In contrast, miR-18a has drawn little attention. No unique function has been ascribed to this miRNA in immune cells, and recently characterized miR-18a–deficient mice did not show any overt immunopathological features (42).

In this study, we uncovered a unique role for miR-18a as a highly inducible inhibitor of Th17 differentiation. Accordingly, miR-18a–deficient mice exhibited increased Th17 responses in airway inflammation models in vivo. We identified Smad4, Hif1a, and Rora as important target genes mediating miR-18a regulation of Th17 cell differentiation.

Mice with loxP sites flanking the miR-17–92 cluster (Mirc1tm1.1Tyj, 008458; The Jackson Laboratory) were crossed to CD4-Cre mice [Tg(CD4-cre)1Cwi; 4196; Taconic Biosciences] to generate T cell–specific miR-17–92–deficient mice. For some experiments, these mice were further crossed with Smad4tm2.1Cxd mice containing loxP sites flanking exon 8 of the Smad4 gene (017462; The Jackson Laboratory) or with mice heterozygous for the spontaneous Rorasg mutation (002651; The Jackson Laboratory) to generate miR-17–92–deficient mice with heterozygous deletion of Smad4 or with one defective Rora allele and appropriate littermate controls. Mice with a targeted deletion of miR-18a (Mir18tm1.1Aven) were described recently (42). Mice with loxP-flanked Dgcr8 alleles (Dgcr8tm1.1Blel) have been described before (43) and were bred to CD4-Cre and R26-stop-EYFP mutant mice [Gt(ROSA)26Sortm3(CAG-EYFP)Hze, 006148; The Jackson Laboratory]. All mice were housed and bred in the specific pathogen–free barrier facilities at the University of California, San Francisco or the Ludwig-Maximilians-Universität München. All experiments were performed according to the Institutional Animal Care and Use Committee guidelines of the University of California, San Francisco or in accordance with the regulations of the Regierung von Oberbayern.

Single-cell suspensions from spleen and lymph nodes were prepared by mincing the tissues between the frosted ends of glass slides. Cells were filtered through fine mesh and counted. CD4+ T cells were enriched with the Easy Sep Mouse CD4+ T Cell Isolation Kit (STEMCELL Technologies). Purified CD4+ T cells were plated at 4 × 106 cells per well in complete medium (RPMI 1640 supplemented with 10% FBS, pyruvate, nonessential amino acids, l-arginine, l-asparagine, l-glutamine, folic acid, 2-ME, penicillin, and streptomycin) in 6-well plates (Corning Costar) or at 1 × 105 cells per well in 96-well, flat-bottom plates (Corning Costar) precoated with 2 μg/ml anti-CD3 (clone 17A2) and anti-CD28 (clone 37.51; both from Bio X Cell). For Th17-polarizing conditions, media were supplemented with anti–IFN-γ (10 μg/ml, clone XMG1.2; Bio X Cell), anti–IL-4 (10 μg/ml, clone 11B11; Bio X Cell), human TGF-β (5 ng/ml; PeproTech), and murine IL-6 (25 ng/ml; PeproTech), unless otherwise stated. In one set of the TGF-β–dosing experiments, no exogenous TGF-β was added to the culture, and cell-derived TGF-β was blocked with anti–TGF-β (20 μg/ml, clone 1D11; Bio X Cell). On day 2 of culture, cells were collected, counted, suspended in transfection buffer together with miRNA mimics, siRNAs, or inhibitors and transfected with the Neon transfection system (Invitrogen). Cells were immediately transferred into fresh culture medium containing Th17-polarizing cytokines plus murine IL-23 (20 ng/ml; R&D Systems) at 4 × 105 cells per well in 96-well flat-bottom plates precoated with anti-CD3 and anti-CD28. Cultured cells were usually analyzed on day 3.5 of initial culture unless otherwise stated.

Cord blood mononuclear cells from anonymous human cord blood donors were isolated by Lymphoprep gradient (1114545; Accurate Chemical & Scientific). CD4+ T cells were isolated from cord blood mononuclear cells using a Dynabeads Untouched Human CD4+ T Cell Isolation Kit (Invitrogen). Cells were stimulated for ∼48 h on plates coated with 2 μg/ml anti-CD3 (clone OKT-3; University of California, San Francisco mAb Core) and 4 μg/ml anti-CD28 (clone 15E8; Miltenyi Biotec) at an initial density of 4–5 × 106 cells per well in complete medium (RPMI 1640 media with 10% FCS, pyruvate, nonessential amino acids, l-arginine, l-asparagine, l-glutamine, folic acid, 2-ME, penicillin, and streptomycin) in six-well plates (Corning Costar). After 2 d of stimulation, cells were collected, counted, suspended in transfection buffer together with miRNA inhibitors, and transfected with the Neon Transfection System (Invitrogen). Cells were immediately transferred into 48-well plates, at a density of 4 × 105 cells per well, precoated with anti-CD3 and anti-CD28 in fresh culture medium containing Th17-polarizing cytokines. For Th17-polarizing conditions, media were supplemented with anti-human IFN-γ (10 μg/ml, clone NIB42; eBioscience), anti-human IL-4 (10 μg/ml, clone MP4-25D2; BioLegend), human TGF-β (5 ng/ml; PeproTech), human IL-6 (25 ng/ml; PeproTech), human IL-1β (20 ng/ml; PeproTech), and human IL-23 (20 ng/ml; PeproTech).

Th17-polarized human or mouse primary CD4+ T cells were transfected with miRNA mimics, small interfering RNAs (siRNAs), or inhibitors at 48 h of cell culture with the Neon Transfection System (Invitrogen), as previously described (41). miRIDIAN microRNA Mimics or siGENOME SMARTpool (both from Dharmacon) were used at 500 nM, and miRCURY LNA microRNA Power family inhibitors (Exiqon) were used at 5 or 20 μM with appropriate controls. MSCV-PGK-hCD25 miRNA sensors for miR-17, miR-18a, miR-92a, and miR-19b (31) were constructed to express EGFP with four perfectly complementary binding sites for the miRNA of interest in the 3′ untranslated region (UTR), as previously described (41). Cells were transduced by spin infection early on day 2 of Th17 cultures and were transfected with miRNA mimics or inhibitors later on day 2 of Th17 cultures. Human CD25+CD4+ T cells were analyzed on day 3.5 for EGFP expression.

3′UTR dual-luciferase plasmids containing near full-length 3′UTRs of Smad4 and Hif1a or an extension 3′UTR for Smad4 were cloned into the psiCHECK-2 luciferase reporter construct (Promega). Primer sequences were Smad4 forward: 5′-TAGTAGCTCGAGCTCTGCAGCTCTTGGATGAA-3′, Smad4 reverse: 5′-TAGTAGGCGGCCGCCATGGGAAAGTCCTGGTAGAG-3′, Hif1a forward: 5′-TAGTAGCTCGAGGGCAGCAGAAACCTACTGCAGG-3′, Hif1a reverse: 5′-TAGTAGGCGGCCGCTAAACGTAAGCGCTGACCCAGG-3′, Smad4 extension forward: 5′-TAGTAGCTCGAGACTGAGTCACTATACGAAGTGG-3′, and Smad4 extension reverse: 5′-TAGTAGGCGGCCGCTTGGCTCTGAAGAGATACTTCC-3′. psiCHECK-2 Rora 3′UTR P1 was described previously (34). CD4+ T cells were transfected on day 2 of Th nonpolarizing (no exogenous cytokines or blocking Abs) culture with luciferase reporter constructs and/or miRNA mimics using the Neon Transfection System. Media were supplemented with 20 U/ml recombinant human IL-2 (National Cancer Institute) on day 2 of culture. Luciferase activity was measured 24 h after transfection with the Dual Luciferase Reporter Assay System (Promega) and a FLUOstar OPTIMA plate reader (BMG Labtech).

Cultured cells were collected, washed, and stained with Abs against cell surface proteins and transcription factors, as described previously (44). Nonspecific binding was blocked with anti-CD16/CD32 (clone 2.4G2; University of California, San Francisco mAb Core), 2% normal rat/mouse serum for mouse T cell staining, or with human FcR binding inhibitor (eBioscience) for human T cells. Dead cells were excluded with Fixable Viability Dye eFluor 780 (eBioscience). For in vitro cell-proliferation experiments, naive CD4+ T cells were labeled with 5 μM CellTrace Violet (Invitrogen), as described previously (44). The proliferation index was calculated using FlowJo software (TreeStar). The Foxp3 staining set (eBioscience) was used for intracellular staining of transcription factors. Intracellular cytokine detection was performed after stimulation with 10 nM PMA and 1 μM ionomycin for 2 h, followed by the addition of 5 μg/ml brefeldin A for another 2 h. Cells were fixed with 4% paraformaldehyde for 8 min at room temperature, washed with ice-cold PBS, and permeabilized with 0.5% saponin. The following fluorochrome-conjugated Abs were used in the study for mouse T cells: anti-CD4 (clones RM4-5 or GK1.5), anti–IL-17A (eBio17B7), anti–IFN-γ (XMG1.2), and anti–IL-13 (eBio13A; all from eBioscience), as well as anti-CD11b (M1/70; BioLegend), anti-CCR6 (140706), anti-RORγt (Q31-378), anti-Siglec F (E50-2440), anti-CD45 (30-F11), and anti-Ly6G (1A8; all from BD Biosciences). Anti-human CD4 (OKT4; BioLegend), anti-human CCR6 (R6H1; eBioscience), and anti-human RORγt (Q21-559; BD Biosciences) were used for human T cells. Apoptotic cells were quantified with the PE Annexin V Apoptosis Detection Kit I (BD Biosciences), according to the manufacturer’s instructions. Samples were acquired on a LSR II or LSRFortessa cytometer (BD Biosciences) and analyzed with FlowJo software (TreeStar).

For the acute 10-d OVA model, sex- and age-matched wild-type (WT) and miR-18∆/∆ mice were sensitized by i.p. injection with 50 μg of OVA (Sigma-Aldrich) in 100 μl of PBS plus 100 μl of Imject Alum Adjuvant (Thermo Scientific). After 7 d, the mice were challenged oropharyngeally with 50 μg of OVA in 20 μl of PBS daily for three consecutive days. Lungs were harvested on day 10 of the experiment. For the 27-d OVA+LPS model, sex- and age-matched WT and miR-18∆/∆ mice were sensitized with 100 μg of EndoFit OVA (InvivoGen) and 10 μg of LPS (Sigma-Aldrich) in 30 μl of PBS delivered by oropharyngeal instillation on days 0, 2, 4, and 11. The mice were then challenged oropharyngeally with 40 μg of OVA in 30 μl of PBS on days 18, 20, 25, and 26. Lungs and lung-draining mediastinal lymph nodes were harvested on day 27. For both airway hypersensitivity models, lungs were digested using the gentleMACS Dissociator (Miltenyi Biotec) and following the lung dissociation kit protocol. Liberase TM was used at 50 μg/ml, and DNase I was used at 25 μg/ml (both from Roche). Cells collected from the lung and lung-draining lymph nodes were analyzed by flow cytometry for surface markers, transcription factors, and cytokines, as described above. Prior to euthanasia, mice were injected retro-orbitally with 2 μg of fluorochrome-conjugated anti-CD45 Ab (clone 30-F11; BD Biosciences) in 200 μl of PBS to distinguish vascular and nonvascular cells in the lung. Routine H&E and periodic acid–Schiff (PAS) staining were performed to assess lung inflammatory cell infiltration and metaplasia of the respiratory epithelium, respectively. A trained histopathologist scored all stained tissue sections.

CD4+ T cells from spleen and lymph nodes of mice were enriched with the EasySep Mouse CD4+ T Cell Isolation Kit (STEMCELL Technologies). Cells were lysed in TRIzol LS (Life Technologies), total RNA was isolated, and RNA was quantified with a ND-1000 spectrophotometer (NanoDrop). Reverse transcription of miRNA was performed with the NCode miRNA First-Strand cDNA Synthesis Kit (Life Technologies). Forward primers were the mature miRNA sequence (45), and a universal reverse primer was used from the kit. Expression values were normalized to 5.8S rRNA (Forward: 5′-ATCGTAGGCACCGCTACGCCTGTCTG-3′). Reverse transcription of mRNA was performed with SuperScript III First-Strand Synthesis for RT-PCR (Invitrogen). Primers for Smad4 total transcript were 5′-CACAATGAGCTTGCATTCCAG-3′ (Forward) and 5′-ACCTTAAACGTCTCTCCTACCT-3′ (Reverse). Primers for Smad4 extended transcript were 5′-CTGAGTCACTATACGAAGTGGAAT-3′ (Forward) and 5′-GTCATTTAGCAGAAGGTGTCTTG-3′ (Reverse). Expression values were normalized to Gapdh (Forward: 5′-GTGTTCCTACCCCCAATGTGT-3′; Reverse: 5′-ATTGTCATACCAGGAAATGAGCTT-3′). Quantitative PCR was performed in technical duplicates using FastStart Universal SYBR Green Master Mix (Roche) on a Realplex2 instrument (Eppendorf).

Excel (Microsoft) and Prism (GraphPad) were used for data analysis. For all figures, bar graphs represent mean + SEM, unless otherwise stated. Z-scores were calculated from mean and SD. *p < 0.05, **p < 0.01, and ***p < 0.001 denote significance. Appropriate statistical analyses were performed for all data and are specified in the figure legends.

The miR-17–92 cluster comprises six miRNAs that can be grouped into four distinct families based on their seed sequence (Fig. 1A). To gain insight into the dynamics of miR-17–92 expression during Th17 cell differentiation, we first cultured purified control (17–92+/+ = CD4-Cre miR-17–92fl/fl) and miR-17–92–deficient (17–92∆/∆ = CD4-Cre+ miR-17–92fl/fl) CD4+ T cells in vitro for 4 d under classical Th17-polarizing conditions (TGF-β+IL-6). We assessed the expression of each miRNA in the miR-17–92 cluster during the course of Th17 differentiation by quantitative PCR (Fig. 1B). All of the miR-17–92 cluster miRNAs were expressed in 17–92+/+ CD4+ T cells under these conditions and, as expected, all six miRNAs were absent in 17–92∆/∆ CD4+ T cells. Although most miRNAs from this cluster were induced upon activation, miR-18a was the most dynamically regulated, showing a >10-fold induction that was sustained over 3 d in culture. Mature miR-92a was not induced together with the other cluster miRNAs. Importantly, representative miRNAs from the paralogous miRNA gene clusters miR-106b–25 and miR-106a–363 (Fig. 1A) were expressed at comparable levels in 17–92+/+ and 17–92∆/∆ CD4+ T cells throughout the time course (Fig. 1C). Consistent with previous reports (46), miR-25 was abundant in developing Th17 cells, whereas miR-363 was barely detectable (Fig. 1C).

FIGURE 1.

miR-18a is active and dynamically regulated during Th17 cell differentiation. (A) Schematic of miRNAs within the miR-17–92 cluster and its paralogous miRNA gene clusters miR-106a–363 and miR-106b–25. Individual miRNA families represented in the three clusters are color coded. (B and C) Naive control (17–92+/+; ○) and miR-17–92-deficient (17–92∆/∆; ▪) CD4+ T cells were cultured in vitro under Th17-polarizing conditions. Cells were harvested at the indicated time points, and quantitative real-time PCR was performed to determine expression levels of individual miRNAs of the miR-17–92 cluster (B) or representative miRNAs from the paralogous miRNA gene clusters miR-106b–25 and miR-106a–363 (C). Expression was normalized to 5.8S rRNA in each sample. (D) 17–92+/+ and 17–92∆/∆ CD4+ T cells were transduced with retroviral sensors expressing EGFP together with four perfectly complementary binding sites for miR-17 (psens17), miR-18a (psens18a), miR-19b (psens19b), or miR-92a (psens92a) in the 3′UTR and analyzed by flow cytometry. Data are pooled from three independent experiments, each with one mouse per genotype (B and C), or are representative of three independent experiments with one mouse per genotype (D). Error bars represent SEM.

FIGURE 1.

miR-18a is active and dynamically regulated during Th17 cell differentiation. (A) Schematic of miRNAs within the miR-17–92 cluster and its paralogous miRNA gene clusters miR-106a–363 and miR-106b–25. Individual miRNA families represented in the three clusters are color coded. (B and C) Naive control (17–92+/+; ○) and miR-17–92-deficient (17–92∆/∆; ▪) CD4+ T cells were cultured in vitro under Th17-polarizing conditions. Cells were harvested at the indicated time points, and quantitative real-time PCR was performed to determine expression levels of individual miRNAs of the miR-17–92 cluster (B) or representative miRNAs from the paralogous miRNA gene clusters miR-106b–25 and miR-106a–363 (C). Expression was normalized to 5.8S rRNA in each sample. (D) 17–92+/+ and 17–92∆/∆ CD4+ T cells were transduced with retroviral sensors expressing EGFP together with four perfectly complementary binding sites for miR-17 (psens17), miR-18a (psens18a), miR-19b (psens19b), or miR-92a (psens92a) in the 3′UTR and analyzed by flow cytometry. Data are pooled from three independent experiments, each with one mouse per genotype (B and C), or are representative of three independent experiments with one mouse per genotype (D). Error bars represent SEM.

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To assess the inhibitory activity of each miRNA family represented within the miR-17–92 cluster, we used previously developed retroviral miRNA activity sensors in developing Th17 cells (31, 41). Each reporter encodes EGFP with four binding sites for the miRNA of interest within an artificial 3′UTR, resulting in EGFP expression that is inhibited by miRNAs of the corresponding family in a sequence-specific fashion (Supplemental Fig. 1). We assessed the activity of the miR-17, miR-18, miR-19, and miR-92 families in 17–92+/+ control and 17–92∆/∆ Th17 cells (Fig. 1D). Sensor EGFP expression was increased in 17–92∆/∆ Th17 cells compared with 17–92+/+ control cells, indicating that endogenous miR-17–92 cluster miRNAs from all four families actively inhibit target mRNAs. However, the increase in EGFP expression was greatest for miR-18 and miR-19 sensors. 17–92∆/∆ Th17 cells displayed only intermediate EGFP expression from miR-17 and miR-92 sensors, suggesting that other family members encoded outside of the miR-17–92 cluster make substantial contributions to miR-17 and miR-92 activity in Th17 cells. These findings were further corroborated by experiments with locked nucleic acid miRNA family inhibitors that are designed to simultaneously inhibit all members of each family (Supplemental Fig. 1). The miR-17 family inhibitor further increased the already elevated expression of the miR-17 sensor EGFP in 17–92∆/∆ Th17 cells. Similar results were obtained for miR-19 and miR-92 family inhibitors and matched sensors, suggesting residual activity beyond what is accounted for by miR-17–92 cluster miRNAs. In contrast, expression of the miR-18 sensor EGFP was unaffected by the miR-18 family inhibitor in 17–92∆/∆ Th17 cells. Taken together, deletion of miR-17–92 was sufficient to nearly abolish miR-18 and strongly reduce miR-19 activity, but other members of the miR-17 and miR-92 families retained substantial activity in 17–92∆/∆ Th17 cells.

To determine the individual contribution of miR-18a and the other miR-17–92 cluster miRNAs to Th17 cell differentiation, we first cultured purified 17–92+/+ and 17–92∆/∆ CD4+ T cells in vitro for 4 d under classical Th17-polarizing conditions (TGF-β+IL-6). 17–92∆/∆ T cells showed increased expression of CCR6 and RORγt (Fig. 2A), with variable IL-17A production (Fig. 2B). To isolate the individual effects of each member of the miR-17–92 cluster on Th17 cell differentiation, miRNA mimics were transfected into activated 17–92∆/∆ CD4+ T cells cultured under Th17-polarizing conditions, and the effect on CCR6, RORγt, and IL-17A expression was assessed by flow cytometry. Strikingly, only miR-18a could significantly rescue the increased frequency of CCR6+RORγt+ cells in 17–92∆/∆ Th17 cells, restoring it to the frequency observed in 17–92+/+ cells (Fig. 2C). In contrast, several other miRNAs of the cluster further increased the already-enhanced frequency of CCR6+RORγt+ cells in 17–92∆/∆ Th17 cell cultures (Fig. 2C). Consistent with a previous report (32), miR-17 family members also enhanced IL-17 production (Supplemental Fig. 2). These data demonstrate that miR-18a exhibits a distinct inhibitory function on CCR6+RORγt+ Th17 cells.

FIGURE 2.

miR-18a inhibits Th17 cell differentiation. Naive 17–92+/+ and 17–92∆/∆ CD4+ T cells were cultured in vitro under Th17 conditions (TGF-β+IL-6) and analyzed on day 3.5 for Th17 marker expression by flow cytometry. (A) Representative contour plots show surface CCR6 and intracellular RORγt costaining of live singlet CD4+ T cells. Frequencies of CCR6+RORγt+ cells, as well as the RORγt gMFI, are quantified in the bar graphs. (B) IL-17A production after restimulation with PMA/ionomycin is shown in a representative histogram, and the frequency of IL-17A+ cells is quantified in the bar graph. (C) Naive 17–92+/+ and 17–92∆/∆ CD4+ T cells were cultured under Th17-polarizing conditions and transfected on day 2 with control miRNA mimics (Ctl mimic) or miRNA mimics of the individual miR-17–92 cluster members. Cells were analyzed on day 3.5 for Th17 marker expression by flow cytometry. The frequencies of CCR6+RORγt+ cells among CD4+ T cells are shown in the bar graph; representative contour plots display surface CCR6 expression and intracellular RORγt costaining of CD4+ T cells. Numbers in quadrants indicate the percentage of CCR6+ and/or RORγt+ live singlet cells. Data are pooled from five (C), or six to eight (A and B) independent experiments, each with one or two mice per genotype. Mean + SEM (A–C). **p < 0.01, ***p < 0.001, two-tailed paired t test with preassigned littermate pairs (A and B) or one-way ANOVA with the Dunnett multiple-comparison posttest comparing all columns with control miRNA mimic-transfected 17–92∆/∆ CD4+ T cells (C).

FIGURE 2.

miR-18a inhibits Th17 cell differentiation. Naive 17–92+/+ and 17–92∆/∆ CD4+ T cells were cultured in vitro under Th17 conditions (TGF-β+IL-6) and analyzed on day 3.5 for Th17 marker expression by flow cytometry. (A) Representative contour plots show surface CCR6 and intracellular RORγt costaining of live singlet CD4+ T cells. Frequencies of CCR6+RORγt+ cells, as well as the RORγt gMFI, are quantified in the bar graphs. (B) IL-17A production after restimulation with PMA/ionomycin is shown in a representative histogram, and the frequency of IL-17A+ cells is quantified in the bar graph. (C) Naive 17–92+/+ and 17–92∆/∆ CD4+ T cells were cultured under Th17-polarizing conditions and transfected on day 2 with control miRNA mimics (Ctl mimic) or miRNA mimics of the individual miR-17–92 cluster members. Cells were analyzed on day 3.5 for Th17 marker expression by flow cytometry. The frequencies of CCR6+RORγt+ cells among CD4+ T cells are shown in the bar graph; representative contour plots display surface CCR6 expression and intracellular RORγt costaining of CD4+ T cells. Numbers in quadrants indicate the percentage of CCR6+ and/or RORγt+ live singlet cells. Data are pooled from five (C), or six to eight (A and B) independent experiments, each with one or two mice per genotype. Mean + SEM (A–C). **p < 0.01, ***p < 0.001, two-tailed paired t test with preassigned littermate pairs (A and B) or one-way ANOVA with the Dunnett multiple-comparison posttest comparing all columns with control miRNA mimic-transfected 17–92∆/∆ CD4+ T cells (C).

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To directly assess the requirement for endogenous miR-18a, we tested Th17 differentiation in miR-18∆/∆ CD4+ T cells from recently developed mice that harbor a targeted deletion of miR-18 while retaining normal expression of all remaining miR-17–92 miRNAs (42). Similar to what we observed for miR-17–92 cluster deletion, loss of miR-18a alone increased the frequency of CCR6+ and CCR6+RORγt+ cells among cultured Th17 cells, with no consistent effect on RORγt expression (Fig. 3A) or IL-17 production (Fig. 3B). These defects could be rescued by transfecting a miR-18a mimic into miR-18∆/∆ CD4+ T cells (Fig. 3C). Because miR-17–92 deficiency considerably impairs CD4+ T cell proliferation, expansion, and survival (21), we next analyzed whether miR-18 deficiency also impacted these processes. Importantly, we did not observe any differences in the proliferative capacity of miR-18∆/∆ CD4+ T cells compared with WT control cells (Fig. 3D). In addition, we did not detect any changes in the rate of apoptotic cells under these culture conditions (Fig. 3E). Taken together, these data demonstrate that miR-18a plays a critical role in limiting Th17 cell differentiation in vitro.

FIGURE 3.

Genetic ablation of miR-18a alone increases the frequency of CCR6+ and CCR6+RORγt+ CD4+ T cells. Naive WT and miR-18∆/∆ CD4+ T cells were cultured in vitro under Th17 conditions (TGF-β+IL-6) and analyzed on day 3.5 for Th17 marker expression by flow cytometry. (A) Representative contour plots show surface CCR6 and intracellular RORγt costaining of live singlet CD4+ T cells. Frequencies of CCR6+ and CCR6+RORγt+ cells, as well as the RORγt gMFI, are quantified in the bar graphs. (B) IL-17A production after restimulation with PMA/ionomycin is shown in a representative histogram, and the frequency of IL-17A+ cells is quantified in the bar graph. (C) Naive WT and miR-18∆/∆ CD4+ T cells were cultured under Th17-polarizing conditions and transfected on day 2 with control miRNA mimics (Ctl mim) or miR-18a mimics. Cells were analyzed by flow cytometry on day 3.5. The frequencies of CCR6+ and CCR6+RORγt+ cells are quantified in the bar graphs. (D) Representative histogram shows CellTrace Violet (CTV)-labeled cells assessed on day 3.5 of culture. Proliferation indices are quantified in the bar graph. (E) Frequencies of early apoptotic (Annexin V+7-AAD) and late apoptotic/dead cells (Annexin V+7-AAD+) among WT and miR-18∆/∆ CD4+ T cells were assessed by flow cytometry on day 3.5 of Th17 culture. Data are pooled from two independent experiments, each with three mice per genotype (A–C) or are representative of two independent experiments with three mice per group (D and E). Mean + SEM. **p < 0.01, ***p < 0.001, two-tailed paired t test with preassigned littermate pairs (A, B, D, and E) or one-way ANOVA with the Dunnett multiple-comparison posttest comparing all columns with control miRNA mimic-transfected miR-18∆/∆ CD4+ T cells (C).

FIGURE 3.

Genetic ablation of miR-18a alone increases the frequency of CCR6+ and CCR6+RORγt+ CD4+ T cells. Naive WT and miR-18∆/∆ CD4+ T cells were cultured in vitro under Th17 conditions (TGF-β+IL-6) and analyzed on day 3.5 for Th17 marker expression by flow cytometry. (A) Representative contour plots show surface CCR6 and intracellular RORγt costaining of live singlet CD4+ T cells. Frequencies of CCR6+ and CCR6+RORγt+ cells, as well as the RORγt gMFI, are quantified in the bar graphs. (B) IL-17A production after restimulation with PMA/ionomycin is shown in a representative histogram, and the frequency of IL-17A+ cells is quantified in the bar graph. (C) Naive WT and miR-18∆/∆ CD4+ T cells were cultured under Th17-polarizing conditions and transfected on day 2 with control miRNA mimics (Ctl mim) or miR-18a mimics. Cells were analyzed by flow cytometry on day 3.5. The frequencies of CCR6+ and CCR6+RORγt+ cells are quantified in the bar graphs. (D) Representative histogram shows CellTrace Violet (CTV)-labeled cells assessed on day 3.5 of culture. Proliferation indices are quantified in the bar graph. (E) Frequencies of early apoptotic (Annexin V+7-AAD) and late apoptotic/dead cells (Annexin V+7-AAD+) among WT and miR-18∆/∆ CD4+ T cells were assessed by flow cytometry on day 3.5 of Th17 culture. Data are pooled from two independent experiments, each with three mice per genotype (A–C) or are representative of two independent experiments with three mice per group (D and E). Mean + SEM. **p < 0.01, ***p < 0.001, two-tailed paired t test with preassigned littermate pairs (A, B, D, and E) or one-way ANOVA with the Dunnett multiple-comparison posttest comparing all columns with control miRNA mimic-transfected miR-18∆/∆ CD4+ T cells (C).

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Next, to test the requirement of endogenous miR-18a for Th17 responses in vivo, we induced asthma-like inflammation in the lungs of control and miR-18∆/∆ mice by sensitizing/challenging these mice with OVA (Fig. 4A). Although we did not observe differences in the numbers of CD4+ T cells or overall CD45+ leukocytes (Fig. 4B), the frequencies of CCR6+ and CCR6+RORγt+ CD4+ T cells were significantly increased in miR-18∆/∆ mice (Fig. 4C). This was also reflected in elevated RORγt geometric mean fluorescence intensity (gMFI) of miR-18∆/∆ CD4+ T cells compared with miR-18+/+ CD4+ T cells. Importantly, frequencies of IL-17–producing cells were also significantly increased among restimulated miR-18–deficient lung CD4+ T cells, whereas frequencies of IL-13–producing Th2 cells and IFN-γ–producing Th1 cells were not affected (Fig. 4D). The number of lung eosinophils, but not neutrophils, was decreased (Fig. 4E), consistent with a shift from Th2-mediated to Th17-mediated inflammation (47) in miR-18∆/∆ airways. We also determined the in vivo function of miR-18 in an inhaled LPS+OVA model of airway inflammation (Fig. 4F). In this model, miR-18 deficiency also increased Th17 cells in the lung without affecting IL-13–producing Th2 cells and IFN-γ–producing Th1 cells (Fig. 4G, 4H). Histologic sections showed dense inflammation surrounding vessels and conducting airways in the lungs of control and miR-18∆/∆ mice (Fig. 4I). However, consistent with a shift from Th2-mediated to Th17-mediated inflammation, miR-18∆/∆ mice showed significantly less mucus-producing lung airway epithelium (Fig. 4J). Finally, miR-18 deficiency also increased Th17 cells in the lung-draining mediastinal lymph nodes (Fig. 4K).

FIGURE 4.

miR-18a deficiency increases lung Th17 cell frequencies in airway inflammation models. (A) Schematic diagram of the 10-d in vivo airway hypersensitivity model with i.p. OVA+alum sensitization on day 0, followed by three consecutive daily challenges with OVA oropharyngeally (o.p.) on days 7, 8, and 9. On day 10, cells from the lungs of WT and miR-18Δ/Δ mice were harvested and analyzed by flow cytometry. (B) Total CD4+ T cell and total CD45+ leukocyte numbers per lung. (C) Frequencies of CCR6+ and CCR6+RORγt+ cells; intracellular RORγt expression levels as determined by gMFI. (D) Frequencies of IL-17A–, IL-13–, or IFN-γ–producing cells among CD4+ T cells after restimulation with PMA/ionomycin. (E) Frequencies of inflammatory cells in the lungs, including eosinophils (CD11b+SiglecF+) and neutrophils (CD11b+Ly6G+). (F) Schematic diagram of the 27-d in vivo airway hypersensitivity model with OVA+LPS sensitizations on days 0, 2, 4, and 11 by oropharyngeal instillation (o.p.), followed by OVA challenge on days 18, 20, 25, and 26. On day 27, cells from the lungs of WT and miR-18∆/∆ mice were harvested and analyzed by flow cytometry. (G) Frequency of CCR6+ and RORγt+ cells among CD4+ T cells. (H) Frequency of IL-17A–, IL-13–, and IFN-γ–producing cells among CD4+ T cells after restimulation with PMA/ionomycin. (I) H&E staining of lungs derived from challenged WT and miR-18∆/∆ mice. Two magnifications are shown. Bronchiole epithelium (Ep) and mixed local inflammation (Infl) are marked for orientation in high-power fields. (J) PAS staining highlighting goblet cell metaplasia of the respiratory epithelium (arrows). Histological scores of PAS staining quantifying mucus-secreting cells are quantified in the bar graph. (K) Lung-draining lymph nodes were analyzed by flow cytometry for the frequency of CCR6+ and RORγt+ cells, as well as for the frequency of IL-17A among CD4+ T cells after restimulation with PMA/ionomycin. Data are pooled from three independent experiments, each with 3–9 mice per genotype [n = 21 mice total per genotype (A–E)] or each with 7–12 mice per genotype [n = 26–31 mice total per genotype (F–H, J, and K)]. Data in (I) are a representative experiment with six mice per genotype. *p < 0.05, **p < 0.01, ***p < 0.001, two-tailed unpaired t test with preassigned littermate pairs.

FIGURE 4.

miR-18a deficiency increases lung Th17 cell frequencies in airway inflammation models. (A) Schematic diagram of the 10-d in vivo airway hypersensitivity model with i.p. OVA+alum sensitization on day 0, followed by three consecutive daily challenges with OVA oropharyngeally (o.p.) on days 7, 8, and 9. On day 10, cells from the lungs of WT and miR-18Δ/Δ mice were harvested and analyzed by flow cytometry. (B) Total CD4+ T cell and total CD45+ leukocyte numbers per lung. (C) Frequencies of CCR6+ and CCR6+RORγt+ cells; intracellular RORγt expression levels as determined by gMFI. (D) Frequencies of IL-17A–, IL-13–, or IFN-γ–producing cells among CD4+ T cells after restimulation with PMA/ionomycin. (E) Frequencies of inflammatory cells in the lungs, including eosinophils (CD11b+SiglecF+) and neutrophils (CD11b+Ly6G+). (F) Schematic diagram of the 27-d in vivo airway hypersensitivity model with OVA+LPS sensitizations on days 0, 2, 4, and 11 by oropharyngeal instillation (o.p.), followed by OVA challenge on days 18, 20, 25, and 26. On day 27, cells from the lungs of WT and miR-18∆/∆ mice were harvested and analyzed by flow cytometry. (G) Frequency of CCR6+ and RORγt+ cells among CD4+ T cells. (H) Frequency of IL-17A–, IL-13–, and IFN-γ–producing cells among CD4+ T cells after restimulation with PMA/ionomycin. (I) H&E staining of lungs derived from challenged WT and miR-18∆/∆ mice. Two magnifications are shown. Bronchiole epithelium (Ep) and mixed local inflammation (Infl) are marked for orientation in high-power fields. (J) PAS staining highlighting goblet cell metaplasia of the respiratory epithelium (arrows). Histological scores of PAS staining quantifying mucus-secreting cells are quantified in the bar graph. (K) Lung-draining lymph nodes were analyzed by flow cytometry for the frequency of CCR6+ and RORγt+ cells, as well as for the frequency of IL-17A among CD4+ T cells after restimulation with PMA/ionomycin. Data are pooled from three independent experiments, each with 3–9 mice per genotype [n = 21 mice total per genotype (A–E)] or each with 7–12 mice per genotype [n = 26–31 mice total per genotype (F–H, J, and K)]. Data in (I) are a representative experiment with six mice per genotype. *p < 0.05, **p < 0.01, ***p < 0.001, two-tailed unpaired t test with preassigned littermate pairs.

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The ability to specifically inhibit miRNAs from the miR-17–92 cluster in a family-wise manner using transfectable inhibitors gave us the opportunity to test whether endogenous miR-18 is important for Th17 differentiation. In line with the observed increase in CCR6+RORγt+ Th17 cell differentiation of miR-18a–deficient cells in vitro (Fig. 3) and in vivo (Fig. 4), inhibition of the miR-18 family significantly increased the expression of CCR6 and RORγt in mouse and human Th17 cells, respectively (Fig. 5). These data indicate that miR-18 acts as an evolutionarily conserved inhibitor of Th17 cell differentiation. Our miRNA family inhibitor experiments also confirmed the miR-17 family’s role in promoting cytokine production (32), because the miR-17 family inhibitor reduced IL-17 production in transfected Th17 cells (Supplemental Fig. 2).

FIGURE 5.

Inhibition of the miR-18 family increases mouse and human Th17 cell differentiation. (A) Murine WT CD4+ T cells polarized under Th17 conditions in vitro and transfected with control inhibitor (Ctl inh.) or miR-18 family inhibitor (18 fam. inh.) on day 2. Cells were analyzed by flow cytometry on day 3.5 of culture. Histograms show representative staining of surface CCR6 and intracellular RORγt expression. Frequencies of CCR6+ cells, as well as RORγt gMFI, are quantified in the dot plots. (B) CD4+ T cells isolated from human cord blood were polarized under Th17 conditions in vitro and transfected with Ctl inh. or 18 fam. inh. on day 2. Cells were analyzed by flow cytometry on day 3.5 of culture. Histograms show representative staining of surface CCR6 and intracellular RORγt expression. Frequencies of CCR6+ cells, as well as RORγt gMFI, are quantified in the dot plots. Dashed lines represent background staining in the CCR6 or RORγt-detecting channels (fluorescence minus one [FMO] controls). Circles (Ctl inh.) and squares (18 fam. inh.) in the dot plots represent the mean of two or three individual transfections for each inhibitor (A and B). Lines connect individual WT mice (A) or individual cord blood donors (B) that received respective inhibitors. Data are pooled from three independent experiments with two or three mice per experiment (n = 7) (A) or from one experiment with three different donor samples (n = 3) (B). *p < 0.05, **p < 0.01, ***p < 0.001, two-tailed paired t test.

FIGURE 5.

Inhibition of the miR-18 family increases mouse and human Th17 cell differentiation. (A) Murine WT CD4+ T cells polarized under Th17 conditions in vitro and transfected with control inhibitor (Ctl inh.) or miR-18 family inhibitor (18 fam. inh.) on day 2. Cells were analyzed by flow cytometry on day 3.5 of culture. Histograms show representative staining of surface CCR6 and intracellular RORγt expression. Frequencies of CCR6+ cells, as well as RORγt gMFI, are quantified in the dot plots. (B) CD4+ T cells isolated from human cord blood were polarized under Th17 conditions in vitro and transfected with Ctl inh. or 18 fam. inh. on day 2. Cells were analyzed by flow cytometry on day 3.5 of culture. Histograms show representative staining of surface CCR6 and intracellular RORγt expression. Frequencies of CCR6+ cells, as well as RORγt gMFI, are quantified in the dot plots. Dashed lines represent background staining in the CCR6 or RORγt-detecting channels (fluorescence minus one [FMO] controls). Circles (Ctl inh.) and squares (18 fam. inh.) in the dot plots represent the mean of two or three individual transfections for each inhibitor (A and B). Lines connect individual WT mice (A) or individual cord blood donors (B) that received respective inhibitors. Data are pooled from three independent experiments with two or three mice per experiment (n = 7) (A) or from one experiment with three different donor samples (n = 3) (B). *p < 0.05, **p < 0.01, ***p < 0.001, two-tailed paired t test.

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To identify relevant miR-18a target genes in Th17 cell differentiation, we compiled a list of 18 previously reported miR-18 family target genes from miRTarBase (48) and performed siRNA-mediated inhibition of these genes in 17–92∆/∆ CD4+ T cells to determine whether any of them are limiting factors for in vitro Th17 cell differentiation and effector cytokine production. siRNA SMARTpool against several of these genes altered the expression of CCR6, RORγt, and/or IL-17 (Supplemental Fig. 3). Following retesting with siRNA SMARTpools to confirm the effects observed in the primary screen (data not shown), the top five target genes whose pooled siRNAs at least partially rescued the increased CCR6 and/or RORγt expression of 17–92∆/∆ CD4+ T cells were examined further. To this end, 17–92∆/∆ CD4+ T cells were separately transfected with three individual unique siRNAs against each gene (Fig. 6A). Multiple siRNAs against Smad4, a transcriptional mediator of TGF-β signaling, reversed the exaggerated increase in CCR6+RORγt+ 17–92∆/∆ T cells. The siRNA SMARTpool against Hif1a, another important transcriptional regulator of Th17 cell differentiation, rescued the increased generation of CCR6+ Th17 cells (Supplemental Fig. 3). Individual siRNAs against Hif1a also partially rescued the increased frequency of CCR6+RORγt+ Th17 cells (Fig. 6A). Targeting either of these genes also reduced IL-17 production (Supplemental Fig. 3). As previously reported (31, 32), siRNAs targeting Pten promoted IL-17A production, but they did so without consistently affecting the generation of CCR6+RORγt+ Th17 cells (Fig. 6A, Supplemental Fig. 3).

FIGURE 6.

Smad4, Hif1a, and Rora are functionally relevant target genes of miR-18a. (A) Naive 17–92+/+ and 17–92∆/∆ CD4+ T cells were cultured under Th17-polarizing conditions and transfected with an siRNA nontargeting control (siNegCtl) or three individual siRNAs against each of five candidate genes. Bars quantify the frequency of CCR6+RORγt+ cells among live CD4+ singlet cells. Dashed line indicates the level of negative-control siRNA-transfected 17–92∆/∆ CD4+ cells. (B) Renilla luciferase activity in 17–92+/+ control and 17–92∆/∆ CD4+ T cells transfected with empty vector or with dual-luciferase reporters for Smad4 3′UTRs assessed 24 h after transfection; results were normalized to firefly luciferase activity and are presented relative to those of transfected 17–92+/+ control CD4+ T cells. (C) Renilla luciferase activity in primary 17–92+/+ control and 17–92∆/∆ CD4+ T cells cotransfected with Smad4 3′UTR luciferase reporters and individual miR-17–92 miRNA mimics or control miRNA mimic (Ctl mimic), assessed 24 h after transfection; results were normalized to firefly luciferase activity and are presented relative to those of Ctl mimic–transfected 17–92∆/∆ CD4+ T cells. (D) Renilla luciferase activity in global miRNA-deficient Dgcr8∆/∆ CD4+ T cells cotransfected with the Smad4 3′UTR extension luciferase reporter and either Ctl mimic or miR-18a mimic (left panel) or with miR-17 mimic (right panel), assessed 24 h after transfection; results were normalized to firefly luciferase activity and are presented relative to those of Ctl mimic–transfected Dgcr8∆/∆ CD4+ T cells. (E) Renilla luciferase activity of 17–92+/+ control and 17–92∆/∆ CD4+ T cells transfected with empty vector or with dual-luciferase reporters for Hif1a 3′UTR regions, as described in (B). (F) Renilla luciferase activity in primary 17–92+/+ control and 17–92∆/∆ CD4+ T cells transfected with Hif1a 3′UTR luciferase reporters together with miR-17–92 miRNA mimics or Ctl mimic, as described in (C). (G) Renilla luciferase activity in primary 17–92+/+ control and miR-17–92-deficient CD4+ T cells transfected with position 1 (P1) of Rora 3′UTR luciferase reporters (see 2Materials and Methods) together with miR-18a mimic or Ctl mimic, as described in (C). (H and I) Frequencies of CCR6+RORγt+ cells from 17–92+/+ control (○), 17–92∆/∆ (▴), and 17–92∆/∆Rorasg/+ (▪) (H) or 17–92∆/∆Smad4∆/+ (▪) (I) cells after 3.5 d of in vitro culture under Th17-polarizing conditions with various doses of TGF-β, as assessed by flow cytometry. Data are pooled from six independent experiments (B and D, left panels) or two to four independent experiments [(C and D, right panels) and (E–I)] with one or two mice per genotype [mean + SEM (A–I)]. *p < 0.05, **p < 0.01, ***p < 0.001, one-tailed paired t test (B, D, and E), one-way ANOVA with the Dunnett posttest [comparing each column with 17–92∆/∆ + Ctl mimic mean (C, F, and G)], and two-way ANOVA with the Bonferroni posttest [comparing 17–92∆/∆ and 17–92∆/∆Rorasg/+ (H) or 17–92∆/∆ and 17–92∆/∆Smad4∆/+ (I)].

FIGURE 6.

Smad4, Hif1a, and Rora are functionally relevant target genes of miR-18a. (A) Naive 17–92+/+ and 17–92∆/∆ CD4+ T cells were cultured under Th17-polarizing conditions and transfected with an siRNA nontargeting control (siNegCtl) or three individual siRNAs against each of five candidate genes. Bars quantify the frequency of CCR6+RORγt+ cells among live CD4+ singlet cells. Dashed line indicates the level of negative-control siRNA-transfected 17–92∆/∆ CD4+ cells. (B) Renilla luciferase activity in 17–92+/+ control and 17–92∆/∆ CD4+ T cells transfected with empty vector or with dual-luciferase reporters for Smad4 3′UTRs assessed 24 h after transfection; results were normalized to firefly luciferase activity and are presented relative to those of transfected 17–92+/+ control CD4+ T cells. (C) Renilla luciferase activity in primary 17–92+/+ control and 17–92∆/∆ CD4+ T cells cotransfected with Smad4 3′UTR luciferase reporters and individual miR-17–92 miRNA mimics or control miRNA mimic (Ctl mimic), assessed 24 h after transfection; results were normalized to firefly luciferase activity and are presented relative to those of Ctl mimic–transfected 17–92∆/∆ CD4+ T cells. (D) Renilla luciferase activity in global miRNA-deficient Dgcr8∆/∆ CD4+ T cells cotransfected with the Smad4 3′UTR extension luciferase reporter and either Ctl mimic or miR-18a mimic (left panel) or with miR-17 mimic (right panel), assessed 24 h after transfection; results were normalized to firefly luciferase activity and are presented relative to those of Ctl mimic–transfected Dgcr8∆/∆ CD4+ T cells. (E) Renilla luciferase activity of 17–92+/+ control and 17–92∆/∆ CD4+ T cells transfected with empty vector or with dual-luciferase reporters for Hif1a 3′UTR regions, as described in (B). (F) Renilla luciferase activity in primary 17–92+/+ control and 17–92∆/∆ CD4+ T cells transfected with Hif1a 3′UTR luciferase reporters together with miR-17–92 miRNA mimics or Ctl mimic, as described in (C). (G) Renilla luciferase activity in primary 17–92+/+ control and miR-17–92-deficient CD4+ T cells transfected with position 1 (P1) of Rora 3′UTR luciferase reporters (see 2Materials and Methods) together with miR-18a mimic or Ctl mimic, as described in (C). (H and I) Frequencies of CCR6+RORγt+ cells from 17–92+/+ control (○), 17–92∆/∆ (▴), and 17–92∆/∆Rorasg/+ (▪) (H) or 17–92∆/∆Smad4∆/+ (▪) (I) cells after 3.5 d of in vitro culture under Th17-polarizing conditions with various doses of TGF-β, as assessed by flow cytometry. Data are pooled from six independent experiments (B and D, left panels) or two to four independent experiments [(C and D, right panels) and (E–I)] with one or two mice per genotype [mean + SEM (A–I)]. *p < 0.05, **p < 0.01, ***p < 0.001, one-tailed paired t test (B, D, and E), one-way ANOVA with the Dunnett posttest [comparing each column with 17–92∆/∆ + Ctl mimic mean (C, F, and G)], and two-way ANOVA with the Bonferroni posttest [comparing 17–92∆/∆ and 17–92∆/∆Rorasg/+ (H) or 17–92∆/∆ and 17–92∆/∆Smad4∆/+ (I)].

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To further validate direct miRNA targeting of candidate target genes from the siRNA screen, we used 3′UTR dual-luciferase reporter assays in primary mouse T cells. Although Smad4 lacks a predicted miR-18 binding site, it had previously been suggested to be a miR-18 target (49). Although we did detect derepression of the annotated Smad4 3′UTR reporter in 17–92∆/∆ CD4+ T cells (Fig. 6B), we could not significantly reverse that effect with any individual miRNA mimic (Fig. 6C). Based on RNA sequencing results (50) (data not shown), we suspected that a longer Smad4 transcript with an extended 3′UTR might be the functionally relevant target in this system. Quantitative RT-PCR confirmed that such a transcript is indeed expressed in naive CD4+ T cells (Supplemental Fig. 4), and a Smad4 3′UTR extension segment luciferase reporter was inhibited by miR-18a and miR-17 in transfected Dgcr8∆/∆ CD4+ T cells (Fig. 6D). The Hif1a 3′UTR reporter was also derepressed in 17–92∆/∆ CD4+ T cells compared with 17–92+/+ CD4+ T cells (Fig. 6E). Importantly, the Hif1a 3′UTR was responsive to miR-18a, as well as miR-17, establishing Hif1a as a novel miR-18 target (Fig. 6F). Finally, miR-18a repressed Rora 3′UTR reporter expression in 17–92∆/∆ CD4+ T cells, providing further evidence (34) that Rora is a direct target of miR-18a, even in the presence of residual miR-17 family activity (Fig. 6G). Biochemical evidence of Ago2 binding further supports direct miR-18a targeting of Smad4, Hif1a, and Rora (51).

These data showed that miR-18a directly targets three transcription factors that are important inducers of the Th17 cell gene-expression program. siRNA experiments also indicated that all three of these transcription factors could be limiting factors for Th17 cell differentiation, suggesting that regulation by miR-17–92 cluster miRNAs could impact their function. To further test this possibility, we generated 17–92∆/∆ CD4+ T cells that were also heterozygous for the Rorasg mutation (which encodes a nonfunctional truncated RORα protein) or a targeted conditional loss-of-function Smad4 allele (Smad4). CD4+ T cells from 17–92∆/∆Rorasg/+ (Fig. 6H) and 17–92∆/∆Smad4∆/+ (Fig. 6I) mice exhibited a decrease in the characteristic elevated frequency of 17–92∆/∆ CCR6+RORγt+ cells to near miR-17–92+/+ control levels, especially when cultured in Th17 conditions with higher doses of TGF-β. Thus, genetically limiting Rora or Smad4 to one allele partially rescued the 17–92∆/∆ phenotype. In summary, these data strongly suggest that Smad4, Hif1a, and Rora are all functionally relevant target genes of miR-18a.

In this article, we showed that miR-18a inhibits Th17 cell differentiation and that this function stands in contrast to other members of the miR-17–92 cluster. Our experiments revealed that miR-18a is the most dynamically regulated miRNA of the miR-17–92 cluster in developing Th17 cells. Individually restoring each miRNA of the miR-17–92 cluster by transfection of 17–92∆/∆ CD4+ T cells with miRNA mimics revealed miR-18a’s ability to inhibit Th17 cell differentiation, whereas other cluster members actually enhanced Th17 cell differentiation. Importantly, these observations were further corroborated by in vitro and in vivo experiments using CD4+ T cells from miR-18a–deficient mice. CCR6 and RORγt expression was also increased in mouse and human CD4+ T cells transfected with miR-18 family–specific inhibitors. Mechanistically, we identified and validated three miR-18a target genes that encode transcription factors important for Th17 cell differentiation, including Smad4, a component of the TGF-β signaling pathway, Hif1a, an important target of miR-210 in its regulation of Th17 cell differentiation (18), and Rora, which we had previously shown to prevent subset-inappropriate gene expression in Tfh cells (34). The finding that miR-18a inhibits Th17 differentiation contrasts strikingly with previous work that described the miR-17–92 cluster’s many roles as a positive regulator of CD4+ T cell differentiation, including Th1 cells (35), Tfh cells (34, 38, 39) and Th2 cells (31). Overall, the current study illustrates how redundancy within families and clusters, coupled with signal-regulated expression, enables miRNAs to confer robustness to T cell differentiation by targeting multiple genes in convergent pathways.

Although Th1 and Th2 cell subsets were initially regarded as stable lineages, more recent work, especially on Th17 and regulatory T cells, has drawn attention to the plasticity and flexibility of Th cell subsets (1, 52). Th cell differentiation is driven by the balanced expression of key transcription factors that form a regulatory network in which small changes in gene expression determine cell fate decisions (52). One important layer of gene regulation that contributes to the control of these processes consists of evolutionarily highly conserved miRNAs (21). For example, we (34) and other investigators (38, 39) have previously shown the importance of the miR-17–92 cluster for promoting robust Tfh cell differentiation. In addition, miR-17–92–deficient Tfh cells inappropriately upregulated a set of genes that are normally associated with Th17 cells, including Ccr6, Il1r1, Il1r2, the cytokine Il22, and the transcription factor Rora, with the last being a direct target of all four miRNA families represented in the miR-17–92 cluster (34). Thus, miR-17–92 emerges as a central regulator of Th cell plasticity (e.g., by preventing a Th17 program in differentiating Tfh cells) (53).

Two previous reports implicated miR-17–92 miRNAs or related family members in the regulation of Th17 cell biology (32, 33). In one study, miR-17 and miR-19b were identified as responsible miR-17–92 miRNAs that promoted Th17-mediated inflammation (32). This is consistent with our finding that miR-17, miR-20a, and miR-19b mimics further enhanced CCR6 and IL-17A expression in 17–92∆/∆ Th17 cells. We extended these findings by showing that inhibition of the entire miR-17 family decreased IL-17A production. Multiple lines of evidence, including new siRNA experiments presented in this article, indicate that Pten is an important miR-17–92 target gene that regulates IL-17 production (31, 32). Nevertheless, we did not observe a consistent decrease in IL-17 production in 17–92∆/∆ CD4+ T cells in vitro, indicating the presence of counterregulation of Th17 differentiation through other miR-17–92 target genes. This possibility was raised in a previous study in which decreased IL-17 production was observed in miR-17–92–deficient CD4+ T cells using different Th17 culture conditions with additional polarizing cytokines (IL-1β and IL-23) and a longer culture period prior to analysis (32). Interestingly, the same study also showed an increase in Rora expression in miR-17–92–deficient CD4+ T cells, consistent with our earlier work on Tfh cells (34) and data presented in the current study showing that Rora is a direct miR-18a target gene involved in Th17 cell differentiation. RORα and RORγt play redundant roles in inducing Th17 signature genes, including Ccr6 (11). A second study showed that miR-20b can directly target Stat3 and Rorc, further suggesting a complex balancing role for the miR-17 family in the regulation of IL-17 production (33). Our results extend these previous and other studies that focused on the various roles of the miR-17–92 cluster in T cell biology and, importantly, identify a novel and distinct role for miR-18a as an inhibitor of Th17 cell differentiation.

The significant impact of miR-18a in determining the net effect of miR-17–92 deficiency may be a product of the differential expression of individual miR-17–92 miRNAs and their related family members in differentiating Th17 cells. T cells strongly express the paralogous miR-106b–25 cluster, which contains two miR-17 family members and one miR-92 family member, and also retain weak, but detectable, expression of the miR-106a–363 cluster. Activity sensors revealed that 17–92∆/∆ CD4+ T cells almost completely lack miR-18 family activity but retain a large fraction of the miR-17 and miR-92 family activity observed in WT CD4+ T cells, as well as a small residual amount of miR-19 family activity. These findings may explain why so many of the requirements for miR-17–92 in T cell biology have been mapped to miR-19 family function.

The importance of miR-18a function in Th17 cells may also relate to its particularly dynamic regulation during Th17 cell differentiation. Mirc1 transcription is induced in activated T cells (54), but expression of the mature miR-17–92 cluster miRNAs is subject to differential regulation. Our data suggest that mature miR-92a induction may be limited by preprocessing of the primary miR-17–92 transcript in developing Th17 cells similarly as in differentiating embryonic stem cells (55). They also indicate further independent regulation of miR-18a processing or stability that maintains it at very low abundance in naive T cells and allows it to be sharply upregulated during T cell activation (Fig. 1B) (46, 56). miR-18a is also the most strongly upregulated miRNA in response to Myc-induced miR-17–92 transcription (57). Previous studies have indicated that miR-17, miR-19a, and miR-20a are also specifically regulated in lymphocytes during disease processes that they promote, including lymphomagenesis and allergic inflammation in asthma (30, 31). Further studies are needed to dissect the intricate regulation of miR-17–92 and its paralogs, as well as the relationships between regulated expression and the biological functions of each mature miRNA. Complete deficiency of miR-17, miR-18, and miR-19 could be studied in mice lacking all three paralogous clusters (39), and each miRNA in the miR-17–92 cluster could be further interrogated using recently reported miR-17–92 allelic series mutant mice (42), which include the miR-18∆/∆ mice that we used in this study.

The distinct regulation and functions of miR-18a and miR-17 illustrate how a cluster of miRNAs can evolve to exert nuanced regulation of gene expression and cell behavior. Phylogenetic comparison suggests that miR-18a was created by a duplication of miR-17 that shifted the miRNA seed sequence by a single nucleotide. As such, miR-18a and miR-17 share many target binding sites, such as in the Smad4 3′UTR extension. Yet these two miRNAs have diverged and acquired independent targets (42), differential regulation of processing and/or stability during T cell activation, and different degrees of redundancy with miRNAs in other clusters. Together, these features allow T cells to translate environmental signals into complex regulation of immune responses through closely related miRNAs within a single miRNA cluster. In future studies, it will be interesting to investigate the upstream mechanisms that regulate the induction of miR-17–92 expression and the differential expression and processing of the individual cluster miRNAs. Given their clear functional differences with regard to regulating Th17 differentiation, this is particularly the case for the closely related miRNAs miR-18a and miR-17.

Our experiments revealed a cell-intrinsic role for miR-18 in Th17 cell differentiation in vitro and corresponding changes in lung inflammation in miR-18∆/∆ mice in vivo. In future studies, it will be important to examine the intrinsic limiting role of miR-18 in Th17 cells in vivo and to explore the interesting possibility that miR-18 may further influence inflammatory responses through independent effects in other cell types. One advantage of using CD4+ T cells from miR-18∆/∆ mice for these studies is that we did not observe any differences in their proliferative or survival capacities. This is in clear contrast to miR-17–92–deficient CD4+ T cells, which show impaired proliferation, expansion, and survival, thus limiting their use for in vivo models in which these altered processes clearly influence the pathophysiological outcomes of the experimental system. In summary, our detailed functional analyses of miR-18a in activated CD4+ T cells in vitro and in vivo highlight the distinct negative impact of this miRNA on Th17 cell differentiation, which is in clear contrast to the function of the other miR-17–92 cluster members. These insights might provide the basis for the development of therapeutic approaches that strengthen the expression of this miRNA or inhibit its target genes.

We thank Sana Patel and Jeanmarie Gonzales for expert technical assistance and Andrea Ventura for providing miR-18a∆/∆ mice.

This work was supported by National Institutes of Health Grants R01HL109102, P01HL107202, U19CA179512, and F31HL131361, a Leukemia & Lymphoma Society scholar award (to K.M.A.), National Institute of General Medical Sciences Medical Scientist Training Program Grant T32GM007618 (to M.M.M.), the National Multiple Sclerosis Society, the University of California, San Francisco Program for Breakthrough Biomedical Research (funded in part by the Sandler Foundation), and Deutsche Forschungsgemeinschaft Grants Emmy Noether Programme BA 5132/1-1 and SFB 1054 Teilprojekt B12 (to D.B.).

The online version of this article contains supplemental material.

Abbreviations used in this article:

gMFI

geometric mean fluorescence intensity

miRNA

microRNA

PAS

periodic acid–Schiff

ROR

RAR-related orphan receptor

siRNA

small interfering RNA

Tfh

T follicular helper

UTR

untranslated region

WT

wild-type.

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The authors have no financial conflicts of interest.

Supplementary data