The IL-4/STAT6 signaling axis establishes a conserved microRNA signature in human and mouse macrophages regulating cell survival via miR-342-3p
- Zsolt Czimmerer†1,
- Tamas Varga†1,
- Mate Kiss†1,
- Cesaré Ovando Vázquez2,
- Quang Minh Doan-Xuan3,
- Dominik Rückerl4,
- Sudhir Gopal Tattikota5,
- Xin Yan5,
- Zsuzsanna S. Nagy1,
- Bence Daniel1, 6,
- Szilard Poliska7,
- Attila Horvath1,
- Gergely Nagy1,
- Eva Varallyay8,
- Matthew N. Poy5,
- Judith E. Allen4,
- Zsolt Bacso3,
- Cei Abreu-Goodger2 and
- Laszlo Nagy1, 6, 9Email author
© The Author(s). 2016
Received: 14 January 2016
Accepted: 10 May 2016
Published: 31 May 2016
IL-4-driven alternative macrophage activation and proliferation are characteristic features of both antihelminthic immune responses and wound healing in contrast to classical macrophage activation, which primarily occurs during inflammatory responses. The signaling pathways defining the genome-wide microRNA expression profile as well as the cellular functions controlled by microRNAs during alternative macrophage activation are largely unknown. Hence, in the current work we examined the regulation and function of IL-4-regulated microRNAs in human and mouse alternative macrophage activation.
We utilized microarray-based microRNA profiling to detect the dynamic expression changes during human monocyte–macrophage differentiation and IL-4-mediated alternative macrophage activation. The expression changes and upstream regulatory pathways of selected microRNAs were further investigated in human and mouse in vitro and in vivo models of alternative macrophage activation by integrating small RNA-seq, ChIP-seq, ChIP-quantitative PCR, and gene expression data. MicroRNA-controlled gene networks and corresponding functions were identified using a combination of transcriptomic, bioinformatic, and functional approaches.
The IL-4-controlled microRNA expression pattern was identified in models of human and mouse alternative macrophage activation. IL-4-dependent induction of miR-342-3p and repression of miR-99b along with miR-125a-5p occurred in both human and murine macrophages in vitro. In addition, a similar expression pattern was observed in peritoneal macrophages of Brugia malayi nematode-implanted mice in vivo. By using IL4Rα- and STAT6-deficient macrophages, we were able to show that IL-4-dependent regulation of miR-342-3p, miR-99b, and miR-125a-5p is mediated by the IL-4Rα–STAT6 signaling pathway. The combination of gene expression studies and chromatin immunoprecipitation experiments demonstrated that both miR-342-3p and its host gene, EVL, are coregulated directly by STAT6. Finally, we found that miR-342-3p is capable of controlling macrophage survival through targeting an anti-apoptotic gene network including Bcl2l1.
Our findings identify a conserved IL-4/STAT6-regulated microRNA signature in alternatively activated human and mouse macrophages. Moreover, our study indicates that miR-342-3p likely plays a pro-apoptotic role in such cells, thereby providing a negative feedback arm to IL-4-dependent macrophage proliferation.
Macrophages display substantial functional heterogeneity, allowing them to participate in diverse aspects of the immune response, including immediate defense against pathogens, regulation of lymphocyte activation, and clearance of cell debris and microbes through phagocytosis, and to contribute to tissue regeneration [1, 2]. Macrophage polarization states and functional properties are determined by the tissue microenvironment containing cytokines, different pathogen-derived molecules, as well as lipid mediators . Two well-established end points of macrophage polarization are classic (M1) and alternative (M2) macrophage activation induced by the Th1-type cytokine interferon gamma and bacterial lipopolysaccharide (LPS) and the Th2-type cytokines interleukin (IL)-4 and IL-13, respectively [3, 4]. M1-type macrophage activation is triggered either by the activation of the Janus kinase (JAK)/signal transducer and activator of transcription (STAT1) axis or the activator protein 1 (AP-1) and nuclear factor kappa-light-chain-enhancer of activated B cells (NFkB) signaling pathways, resulting in enhanced bactericidal capacity and pro-inflammatory properties [3, 5]. In contrast, IL-4 activates the IL4Rα/JAK/STAT6 and phosphoinositide 3-kinase (PI3K) pathways, both of which contribute to alternative macrophage activation . M2-type macrophages possess a characteristic gene expression signature endowing them with anti-inflammatory and immune regulatory properties. The significance of alternative macrophage activation has been described in a range of physiological and pathological processes, including hypersensitivity, anti-helminthic immune responses, fibrosis, sepsis, and tumor progression [4, 6–8]. The functional properties associated with distinct macrophage activation states require tight but plastic regulation of activation-specific gene expression programs at the transcriptional and post-transcriptional levels . In the past decade it has been shown that microRNAs (miRNAs) are important components of post-transcriptional fine tuning of gene expression in mammals [10–12].
miRNAs are short, 18–25-nucleotide-long, single-stranded, non-coding RNA molecules. They are transcribed from different regions of the genome, including intergenic and intronic/exonic regions of protein-coding genes, by RNA polymerase II. Primary transcripts are processed in two steps during miRNA biogenesis by the RNase III enzymes Drosha and Dicer . The mature miRNAs are incorporated into the RNA-induced silencing complex (RISC)  and generally bind the 3′ untranslated region (3′ UTR) of target messenger RNAs (mRNAs) and act as negative regulators of gene expression through inhibition of protein synthesis and/or induction of target mRNA degradation [10, 11]. Several miRNAs, such as miR-155, miR-21, and miR-146, are induced in macrophages in response to LPS, suggesting that they have a role in the regulation of macrophage activation and M1-type polarization [14–18]. In addition, a number of miRNAs have recently been linked to alternative macrophage activation. For instance, miR-125b-5p, miR-199b, and miR-378-3p showed elevated expression in nematode infection-elicited alternative macrophage activation . Among these miRNAs, miR-378-3p has been shown to repress IL-4-driven macrophage proliferation through the modulation of the PI3K/AKT signaling pathway . Furthermore, miR-124, miR-324-5p, and miR-511-3p have been shown to fine-tune macrophage functions during allergic inflammation and tumor development, conditions characterized by the predominance of alternative macrophage activation [20–23]. Finally, miR-193b has been found to be induced by IL-4 in human macrophages, although its function remains unknown .
Despite the fact that IL-4-induced alternative macrophage activation-specific miRNAs have been identified in human and mouse in in vitro as well as in vivo models [19, 24], the regulatory mechanisms of miRNA expression and their role during alternative macrophage activation remain poorly understood. In this work we identified the human alternative macrophage activation-specific microRNome using miRNA microarray and quantitative PCR (qPCR) techniques. We found IL-4-dependent induction of miR-342-3p and miR-193b and repression of miR-99b and miR-125a-5p. Intriguingly, IL-4-dependent regulation of miR-342-3p, miR-99b, and miR-125a-5p is conserved between human cells and mouse bone marrow-derived macrophages. Furthermore, these miRNAs show the same expression pattern in an in vivo mouse model of parasitic infection. The IL-4-regulated expression of miR-342-3p, miR-99b, and miR-125a-5p is dependent on IL-4Rα and STAT6 expression as demonstrated by loss of function genetic experiments. Additionally, miR-342-3p and its host gene, Evl (Ena/Vasp-like), are co-regulated during IL-4-induced alternative macrophage activation in mouse. Chromatin immunopreciptiation (ChIP) confirmed that STAT6 directly binds to the regulatory region of the EVL gene in IL-4-stimulated human and mouse macrophages. Functional analyses combined with miRNA manipulation studies demonstrated that miR-342-3p decreases the viability of macrophages by inducing apoptosis. In silico target prediction and functional analyses identified the anti-apoptotic direct target gene network of miR-342-3p, which includes Bcl2l1. These findings suggest that enhanced miR-342-3p expression is a component of a negative feedback loop controlling excessive macrophage proliferation during Th2-type inflammatory responses.
Cell isolation and culture
Isolation of human monocytes, mouse bone marrow cells, and mouse peritoneal macrophages was done as described earlier with minor modifications [19, 25, 26]. CD14+ human monocytes were isolated from platelet-free buffy coats from healthy donors by Ficoll gradient centrifugation followed by immunomagnetic cell separation with anti-CD14-conjugated microbeads (VarioMACS Separation System, Miltenyi Biotec.). Monocytes were cultured in multiwell culture plates in RPMI 1640 (Sigma-Aldrich) supplemented with 10 % fetal bovine serum (Invitrogen) and penicillin/streptomycin (Sigma-Aldrich). For macrophage differentiation and alternative macrophage activation, freshly plated monocytes were treated with IL-4 (100 ng/ml; PreproTech). Cells were harvested 12 and 72 h after cytokine treatment. Bone marrow isolation and bone marrow-derived macrophage (BMDM) differentiation were performed as described before . Bone marrow was flushed from the femur of wild-type (WT), IL-4Rα, and STAT6 knockout (KO) male animals. Cells were purified through a Ficoll-Paque gradient (Amersham Biosciences, Arlington Heights, IL, USA) and cultured in DMEM containing 20 % endotoxin-reduced fetal bovine serum and 30 % L929 conditioned medium for 5 days. For alternative macrophage activation we treated freshly isolated bone marrow cells with IL-4 (5 ng/ml; PreproTech). Culture medium was changed for Macrophage SFM medium (Gibco) on the 5th day. IL-4 (20 ng/ml) was further added to the culture of macrophage colony stimulating factor (MCSF)- and IL-4-pretreated macrophages for 24 h for the alternative macrophage activation. Peritoneal exudate cells from Brugia malayi-infected animals were seeded at 5 × 106 cells per well to six-well cell culture plates (NUNC) in RPMI, 5 % fetal calf serum, 2 mM L-glutamine, 0.25 U/mL penicillin, and 100 mg/mL streptomycin. After 4 h of incubation at 37 °C, 5 % CO2, non-adherent cells were washed off and the adherent cells were lysed in 700 μL Qiazol.
Animals and infection
Wild-type mice (WT; C57BL/6J or BALB/c), Stat6-deficient (Stat6 KO) mice on C57BL/6 J, and IL-4 receptor-α-deficient (IL-4Rα KO) mice on a BALB/c background were housed under minimal disease conditions. Surgical implantation of adult B. malayi parasites into WT (C57BL/6) mice was carried out as described previously  and performed by the same group and at the same facilities as in case of of the results presented by Jenkins et al. .
Macrophages were resuspended in staining medium (phenol-red free DMEM, 10 mM HEPES, 2 % fetal bovine serum) and incubated with anti-mouse CD206 (AbD Serotec), anti-human CD206 (BD Biosciences), or corresponding unspecific isotype control antibodies for 20 minutes at 4 °C. Cells were washed and resuspended in staining medium for flow cytometry. Data acquisition was performed using a FACSAria III instrument (BD Biosciences) and data were analyzed with FlowJo software.
Arginase activity measurement was performed as previously described with minor modifications .
miRNA microarray and data analysis
Global miRNA expression data were obtained with Affymetrix miRNA 1.0 arrays (Affymetrix). Total RNA samples were labeled using an Affymetrix FlashTag Biotin HSR RNA Labeling Kit according to the manufacturer’s protocol. Briefly, 500 ng of total RNA samples were poly(A)-tailed using poly A polymerase enzyme and ATP at 37 °C for 15 minutes, then biotinylated by ligating biotin-labeled fragment to the 3′ end. Labeled samples were hybridized on miRNA 1.0 arrays at 48 °C and 60 rpm for 16 h. After that, arrays were washed and stained using the standard Affymetrix protocol with an Affymetrix Hybridization, Wash, and Stain Kit on a FS 450 fluidic station instrument. The arrays were scanned using an Affymetrix GeneChip Scanner 3000 7G instrument.
The miRNA CEL files were processed with R, using affy and mirna10cdf packages [29, 30]. Raw expression values were processed with the robust multi-array average (RMA) procedure (background adjustment, quantile normalization, and summarization of log-expression values for each miRNA for each array) . Only probes annotated as human miRNAs were kept. Lowly expressed probes were removed if expression was lower than the median in at least two samples. The interquartile range was calculated across all samples for each probe to determine the most variable probe of those mapped to the same miRNA; 563 probes were annotated, expressed, and unique. Quality control was performed using principal component analysis. Differential expression analysis was performed using the limma package [32, 33] by fitting a linear model for each probe then using an empirical Bayes method to get moderated t-statistics. The Benjamini and Hochberg method was used to calculate false discovery rates (FDRs) .
Small RNA-Seq and data analysis
A cDNA library for small RNA-Seq was generated from 1 μg total RNA using an Illumina TruSeq Small RNA Sample Preparation kit, according to manufacturer’s protocol. Briefly, after ligation of 3′ and 5′ RNA adapters, reverse transcription was performed to synthesize cDNA; then the cDNA was amplified using a common primer and primer-containing index sequence. The amplified product/library was excised from gel (6 % Novex TBE PAGE gel) and, after purification, the libraries were quantified by Qubit fluorometer and checked on a BioAnalyzer 2100 using DNA1000 chip (Agilent Technologies). The small RNA-Seq libraries were diluted to 10 nM and five libraries were pooled together before sequencing. A single-read 50-bp sequencing run was performed on an Illumina HiScan SQ instrument (Illumina, San Diego, CA, USA). Each library pool were sequenced in one lane of a sequencing flow cell; 16–18 million reads per sample was obtained. Sequencing results from small RNA libraries were 3′ adapter trimmed and de-duplicated using the reaper and tally command-line tools from Kraken . The processed small RNA-Seq reads were mapped to all available Mus musculus mature miRNA sequences (miRBase v21 ). Read mapping was performed using Bowtie . The mapping process for reads longer than ten nucleotides allowed two mismatches. Quantification of mapped reads with minimum and maximum lengths of 18 and 24 nucleotides overlapping mature miRNA regions (miRBase v21) was performed using the R package GenomicRanges [38, 39]. The minimum overlap between mapped reads and mature regions was 16. We filtered out mature miRNAs with counts per million (cpm) lower than or equal to 4 cpm across at least three libraries. Differential expression analysis was performed using the edgeR package  fitting a negative binomial generalized linear model for each mature miRNA and then the likelihood ratio test was performed. The comparisons were: WT in the presence of IL-4 versus WT control, WT in the presence of IL-4 versus STAT6 KO in the presence of IL-4, STAT6 KO in the presence of IL-4 versus STAT6 KO control, and WT control versus STAT6 KO control. The Benajmini and Hochberg method was used to calculate FDRs .
Total RNA was isolated from cells using Tri Reagent (MRC) according to the manufacturer’s protocol. For quantification of mRNAs, pri-miRNAs, and mature miRNAs, reverse transcription was performed using a High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems). Reverse transcription (RT) primers for mature miRNAs were supplied by Applied Biosystems. Transcript quantification was performed by quantitative real-time RT-PCR using Taqman probes or SYBR Green assays (selfmade assays) and Taqman Gene Expression Assays (Applied Biosystems).
For semi-quantitative PCR-based quantification of miR-125a_Ncrna00035 in unstimulated and IL-4-stimulated macrophages, specific primers were designed with Primer3. The PCR program used for semi-quantitative amplification was: 95 °C for 10 minutes, followed by 25 cycles of 30 s at 95 °C, 60 s at 65 °C, and 30 s at 72 °C. PCR products were separated by agarose gel electrophoresis and visualized by GelRed staining. qPCR assays and primer sequences are listed in Additional file 1.
GRO-seq and ChIP-seq data analysis
All samples were aligned to the mouse reference genome (mm9)  and RNA-Seq data were analyzed using the standard TopHat/Cufflinks pipeline . For ChIP-seq and GRO-seq (global run-on sequencing) data an integrative analysis pipeline was applied . Data were visualized in Integrated Genome Browser .
ChIP was performed as previously described with minor modifications . Briefly, cells were crosslinked with DSG (Sigma) for 30 minutes and then with formaldehyde (Sigma) for 10 minutes. After fixation, chromatin was sonicated with a Diagenode Bioruptor to generate 200–1000-bp fragments. Chromatin was immunoprecipitated with antibodies against pre-immune IgG (Millipore, 12–370), H3K27ac (Abcam, ab4729) and STAT6 (Santa Cruz, sc-981). Chromatin–antibody complexes were precipitated with Protein A-coated paramagnetic beads (Life technologies). After six washing steps, complexes were eluted and reverse crosslinked. DNA fragments were column purified (Qiagen, MinElute). The amount of immunoprecipitated DNA was quantified with a Qubit fluorometer (Invitrogen). DNA was applied for qPCR analysis. Primer sequences are listed in Additional file 2.
To determine the potential function of miR-342-3p, RAW264.7 cells were transfected with 30 nM miR-342-3p precursor (Applied Biosystems) or scrambled miRNA negative control (Applied Biosystems) using DharmaFECT 3 (Thermo Scientific) in 12- and 96-well plates (Sigma-Aldrich) and eight-well chamber slides (Thermo Scientific).
mRNA microarray and computational target identification
The raw sample CEL files were processed within R using the affy, org.Mm.eg.db and mogene10stv1cdf packages [29, 46, 47] Similarly to the miRNA microarray processing, raw expression values were processed with the RMA procedure . The interquartile range was calculated across all samples for each probe in an identical way to the miRNA microarray methodology to remove probes mapping to the same transcript. Quality control was performed using principal component analysis. Differential expression analysis was performed as mentioned in the “miRNA microarray and data analysis” section, using the limma package  to fit a linear model to each probe then using an empirical Bayes method to determine moderate t-statistics. The comparison used was transfection with miR-342-3p versus miR-negative control miRNA mimics in the mouse macrophage cell line RAW264.7. The Benjamini and Hochberg method was used to calculate FDRs . The comparison was used to verify direct effects due to miR-342-3p regulation. Sylamer  was used to quantify the enrichment or depletion of miRNA seed matches in the 3′ UTRs of genes. The genes with enriched 3'UTRs were ordered according to the t-statistic (differential expression). Prediction of functional miRNA targets was performed by TargetExpress (C Ovando-Vázquez, D Lepe-Soltero, C Abreu-Goodger, submitted). TargetExpress is a support vector machine combination based on TargetScan  and microT-CDs  predictions and an expression profile. In this case, the miR-negative control expression profile was included in the TargetExpress model.
The 3′ UTR sequences corresponding to selected downregulated and computationally predicted anti-apoptotic mmu-miR-342-3p target genes were obtained using the TxDb.Mmusculus.UCSC.mm10.ensGene and GenomicFeature packages [38, 51] Conservation scores for these 3′ UTR sequences were obtained from the UCSC GenomeBrowser using the rtracklayer package , selecting the track cons60way to obtain the phastCons60wayPlacental table.
Cell number analysis
Propidium iodide (PI)-based cell number analysis was performed as previously described with minor modifications . Briefly, macrophages attached to the bottom of 96-well plates were permeabilized by Triton X-100, stained by PI and measured in a Synergy HT microplate reader (Bio-Tek Instruments, Winooski, VT, USA) at 530/25 nm excitation and 645/40 nm emission. Cell numbers were determined using the cell number reference curve.
Cell viability analysis
Resazurin-based viability staining (Promega) was carried out according to the manufacturer’s protocol. Neutral red staining was performed as previously described with minor modifications .
Cell cycle analysis
We plated 2 × 104 cells per well in Labtek 8-well chambered coverslips and transfected them with appropriate miRNAs. Cell fixation and PI staining were performed 48 h post-transfection as previously described  with minor modifications. Raw264.7 cells were fixed on chamber slides using −20 °C methanol at −20 °C overnight. Fixed cells were covered with 5 μg/ml PI (Sigma-Aldrich) and 200 μg/ml RNase A (Sigma-Aldrich) in 0.4 ml phosphate-buffered saline for 2 h at room temperature in the dark. Measurements were performed on a slide-based iCys Research Imaging Cytometer (CompuCyte Corporation, Westwood, MA, USA). The DNA dye was excited by the 488 nm laser line and emission was collected in the red channel with a 650 nm longpass filter on a linear scale. Single cells were recognized according to their size and circularity and cells were further gated according to their integral fluorescence intensity and maximum pixel intensity on a two-dimensioanl scattergram of PI signal. Data from the one-dimensional histogram of the DNA staining was exported and fitted with the automatic one cycle diploid model of the Modfit LT 3.0 software (Verity Software House) with AutoDebris compensation, AutoAggregate Compensation, and Apoptosis Model. In the measurements, the G1–G2 linearity ratio was around 1.8 and the RCS (Reduced Chi-Square, a measure of goodness of fit) was less than 5. Measurements were repeated three times and approximately 1000–2000 cells were collected from each well.
Raw264.7 cells were plated and treated in a similar manner as for the cell cycle analysis. Annexin V-FITC/PI staining was performed as previously described  with minor modifications. Forty-eight hours following miRNA transfection, the culture medium was replaced by 100 μl AB buffer (140 mM NaCl and 2.5 mM CaCl2), 10 μl Hoechst 33342 solution (50 μg/ml stock), and 10 μl propidium iodide solution (50 μg/ml stock) and 5 μl FITC-conjugated annexin V (AV) was added (according to the description of the MBL Apoptosis Detection Kit) for 15 minutes at 37 °C. Laser-scanning cytometric measurements were made using a iCys Research Imaging Cytometer (CompuCyte Corporation, Westwood, MA, USA). Hoechst, which stains all cells, was excited at 405 nm and detected with a 463/20 bandpass filter; PI, which stains dead cells, was excited at 488 nm and detected with a 650 nm longpass filter; and the AV signal, which indicates apoptotic cells, was excited at 488 nm and detected with a 530/30 bandpass filter. Single cells were selected by size and circularity and additionally gated by their DNA content. Early apoptotic AV+/PI−, late apoptotic AV+/PI+, and unapototic AV−/PI− cells were detected using a quadrant gate in the PI versus AV dot plot with respect to unlabeled and miRNA untreated cells.
PsiCHECK2 dual luciferase vector (Promega) was used to confirm the function of the putative miR-342-3p binding sites in the Bcl2l1 3′ UTR. For luciferase reporter assays, 320 bp (miR-342-3p_1) and 309 bp (miR-342-3p_2) of the 3′ UTR of the Bcl2l1 gene, including the mir-342-3p target sites, were amplified by PCR using F1/R1 and F3/R3 primer pairs with XhoI and NotI sites. PCR was performed on mouse macrophage-derived genomic DNA. The XhoI/NotI-digested PCR product was cloned into the XhoI/NotI-digested psiCHECK2 dual luciferase vector. F1/R2 and F2/R1 as well as F3/R4 and F4/R3 primers were used to delete the mir-342-3p target sites from the 3′ UTR. After mixing the two PCR products and digestion with XhoI and NotI, the 3′ UTR fragment with deleted mir-342-3p binding sites was cloned into a XhoI/NotI-digested psiCHECK2 vector. Primer sequences are given in Additional file 2.
Luciferase assays were performed as described previously . Brifely, HEK293 cells were grown on 24-well plates to 70–80 % confluence. Transfection was performed using PEI with 0.1 μg of pRL-TK (Rr-luc) containing 3′ UTR sequences and 0.1 μg of pGL3 control vector (Pp-luc) (Promega) for either a specific miR-342 or small interfering RNA control. Cell lysates were collected 30–36 h post-transfection and luciferase activity was determined using a Dual-Luciferase® Reporter Assay System (Promega, Mannheim, Germany). Renilla luciferase activity was normalized to firefly luciferase activity for each cell culture well and averaged across six well repetitions per condition.
The two-tailed Student’s t-test was used to evaluate the significance of the differences between two groups; p values <0.05 were considered significant.
Identification of the human alternatively activated macrophage-specific miRNA signature
Conserved IL-4Rα/STAT6 signaling-dependent regulation of miR-342-3p, miR-99b, and miR-125a-5p expression during in vitro and in vivo mouse alternative macrophage activation
Direct Stat6-dependent induction of miR-342-3p and its host gene, EVL, during alternative activation of murine and human macrophages
Intronic miRNAs and their host genes are often co-regulated, although there are known instances when intronic miRNAs are regulated by their own promoter region independently of that of the host gene [61, 62]. miR-342-3p is encoded within the third intron of the EVL gene in humans and mice and its expression showed coordinated regulation in both human colorectal cancer and multiple myeloma [63, 64]. Therefore, we decided to further characterize the potential regulatory mechanism controlling the expression of the host gene and the miRNA during human and mouse alternative macrophage activation. For this reason we analyzed publicly available GRO-seq data from unstimulated and ChIP-seq datasets from both IL-4-stimulated and unstimulated mouse macrophages [65, 66].
Moreover, we examined the macrophage lineage-specific transcription factor Pu.1 and IL-4-activated STAT6 binding at the predicted EVL-associated functional domain/chromatin loop. First, CTCF and Rad21 ChIP-seq data sets from unstimulated macrophages were used for the functional domain/chromatin loop prediction. Functional chromatin domains were defined using the following criteria: (i) common CTCF and Rad21 binding at the starting points of chromatin interactions; and (ii) a predominantly convergent direction of the CTCF elements under CTCF/Rad21 co-peaks [67–70]. One distinct Evl-associated putative functional domain (loop) was predicted using these features (Fig. 4a). Next, Pu.1-bound genomic regions were identified and examined within the predicted functional domain in both unstimulated and IL-4-treated macrophages but Pu.1-binding was not influenced by IL-4 stimulation (Fig. 4a). In addition, IL-4-induced Stat6 peaks were detected both upstream and downstream of the TSS of the Evl gene in mouse macrophages  (Fig. 4a).
Next we validated two Stat6-bound genomic regions −4 kb (mEvl_Stat6 site A) and +4 kb (mEVL_Stat6 site B) from the TSS. ChIP followed by qPCR showed an enrichment of Stat6 binding at these sites in IL-4-stimulated WT macrophages relative to untreated WT and IL-4-treated Stat6-deficient macrophages as well as IgG controls (Fig. 4d). These are the likely enhancers controlling Evl as well as miR-342-3p induction. In order to determine whether IL-4-mediated co-regulation of EVL and miR-342-3p is conserved between human and mouse, we measured their expression in human monocyte-derived unstimulated and IL-4-stimulated macrophages. As shown in Fig. 4e and Additional file 6b, IL-4 induced EVL expression in all three donors after 12 h, which then was reduced at later time points. miR-342-3p followed similar but delayed kinetics upon IL-4 treatment as its elevated expression was only detectable after 24 h (Fig. 4f; Additional file 6c). In order to assess whether STAT6 binds the genomic locus of human EVL, we utilized public human STAT6 ChIP-seq datasets from IL-4 stimulated Th2-type T cells . Interestingly, a genomic region within +0.3 kb of the TSS of the EVL gene was occupied by STAT6 in these cells. Therefore, we hypothesized that this region might also be occupied by STAT6 in human alternatively activated macrophages. This could be confirmed by ChIP-qPCR in two independent donors (D1 and D2; Fig. 4g). These findings suggest that miR-342-3p and its host gene, EVL, are coordinately regulated by IL-4 via direct DNA binding of STAT6 in both mouse and human macrophages.
Alternative macrophage activation-specific repression of the miR-99b-125a miRNA polycistron is mediated at the transcriptional level by IL-4/STAT6 signaling
However, Pu.1- and Stat6-binding activity could not be detected within the predicted functional domain of pri-miR-99b-125a. These findings suggest that although the IL-4-mediated regulation of miR-99b and miR-125a expression is STAT6-dependent, (i) it might not be regulated via direct STAT6 DNA binding or (ii) the regulatory region might be outside of that explored within this study (Fig. 5a). Interestingly, H3K27 acetylation (the active histone mark) of the TSS of pri-miR-99b-125a was IL4-dependently reduced in WT but not in STAT6-deficient BMDMs (Fig. 5a, d), which further suggests that IL-4 represses transcription of the miR-99b-125a polycistron in a STAT6-dependent manner.
In addition, we found nascent RNA transcription at the genomic locus of the neighboring Spaca6 gene, although no H3K4m3 peak was detected at the 5′ end of this gene (Fig. 5a). Based on this observation, we hypothesized that the miR-99b-125a polycistron and Spaca6 might have common primary transcripts in macrophages. In order to test this, we used RT-PCR assays in which the PCR primers detected both miR-125a and the first predicted exon of Spaca6 (the genomic localization of a common pri-miR-99b-125a and Spaca6-specific primer pair is shown in Additional file 9). As shown in Fig. 5e, the expected 440-nucleotide PCR product was present in untreated macrophages and IL-4 treatment reduced the expression of this transcript (Fig. 5e). These findings suggest that the miR-99b-125a polycistron and Spaca6 form one STAT6-regulated transcription unit and they are encoded by the same primary transcript.
miR-342-3p regulates cell proliferation and apoptosis-associated signaling pathways at the post-transcriptional level in macrophages
miR-342-3p acts as a regulator of macrophage cell number via reduction of cell viability and induction of apoptosis
Our transcriptome analysis revealed cellular proliferation and cell death as major target processes regulated by miR-342-3p. Proliferation of local macrophage population is induced by IL-4-mediated Th2-type inflammation . Furthermore miR-342-3p has been described to be aberrantly downregulated in different malignancies, including colorectal and breast cancers [58, 63]. In addition, overexpression of miR-342-3p was found to induce apoptosis and block cancer cell proliferation [59, 63]. Thus, we hypothesized that IL-4-induced miR-342-3p expression could be a potential negative feedback mechanism controlling excessive macrophage expansion upon Th2-type cytokine stimulus.
Many Th1- and Th2-type inflammation-linked miRNAs, including miR-155, miR-146, and miR-324-5p, appear to be conserved in both mouse and human macrophages and are induced by LPS or IL-4, suggesting that the regulation of miRNA expression and function during classic or alternative macrophage activation is conserved [15, 18, 20, 24, 75].
Our data presented here suggest that additional conserved miRNAs exist in alternatively activated macrophages. We studied the regulation of miR-342-3p, miR-99b, and miR-125a-5p expression during IL-4-induced differentiation in vitro in human and murine macrophages as well as in nematode implantation-induced alternatively activated murine macrophages in vivo. We found that macrophage-specific, IL-4-dependent regulation of these miRNAs showed conserved regulation in human and mouse. It has been previously shown that these miRNAs are expressed in myeloid cells and participate in the regulation of macrophage phenotype, but their function in alternative macrophage activation has not been described before [24, 76–80].
miR-125a and miR-125b, containing the same seed sequences, are induced during classical macrophage activation and potentiate M1 polarization through direct inhibition of anti-inflammatory Tnfaip3 (A20) and Irf4 expression [24, 81, 82]. Intriguingly, we found that miR-125a expression is reduced during alternative macrophage activation both in vitro and in vivo. It has been previously described that the common miR-125a/miR-125b target gene Irf4 facilitates M2 macrophage polarization and host response against helminth infection [83, 84]. In addition, several target prediction algorithms predict Irf4 as a direct target gene of miR-99b. These data raise the possibility that decreased miR-125a and miR-99b levels might contribute to alternative macrophage activation by leading to increased Irf4 expression. However, miR-125b shows opposite regulation during in vitro and in vivo alternative macrophage activation, though regulation of miR-125b expression was not studied at early time points of nematode infection-induced M2 macrophage activation . These findings collectively suggest a complex role of miR99b, miR-125a, and miR-125b in the regulation of alternative macrophage activation. Nevertheless, further studies are needed to explore the time-dependent regulation and dissect the exact roles of these miRNAs during M2 polarization.
IL-4 stimulation leads to the activation of both the JAK/STAT6 and PI3K signaling pathways in myeloid cells . However, it was previously unknown which pathways play critical roles in the regulation of the macrophage microRNome during IL-4-dependent alternative macrophage activation. We observed that the majority of IL-4-regulated miRNAs were strictly STAT6-dependent in mouse macrophages, including miR-342-3p, miR-125a-5p, and miR-99b-5p, as well as the previously studied miR-511-5p and miR-324-5p. In addition, genome-wide mapping of Stat6-binding in IL-4-stimulated T cells showed that activated Stat6 binds intronic and exonic regions as well as distant enhancer regions of its target genes [71, 85]. Accordingly, we found that IL-4 induces Stat6 binding of an adjacent genomic region of Evl in mouse and human macrophages (+4 kb and −4 kb in mouse; +0.3 kb in human), suggesting that Evl and miR-342 are direct targets of IL-4/Stat6 signaling in both human and murine alternative macrophage activation. Interestingly, elevated expression of miR-342-5p (derived from the 5′ strand of the pre-miR-342 miRNA precursor) in inflammatory macrophages has been linked to the formation of atherosclerotic lesions . Both these results and our findings suggest that pri-miRNA expression and miRNA strand selection may be regulated in macrophages by different external stimuli. Interestingly, 11 miRNAs showed IL-4-dependent regulation in STAT6 deficient macrophages, suggesting the participation of STAT6-independent signaling pathway(s) in the regulation of these miRNAs.
Rapid macrophage accumulation is observed in different pathological conditions, including pathogen infections and inflammation . Although the general paradigm of macrophage accumulation is that monocytes or other hematopoietic cells are recruited to sites of inflammation and differentiate into macrophages [87, 88], the rapid proliferation of resident macrophages in adipose tissue of obese animals and atherosclerotic lesions was also described recently [89, 90]. In addition, local IL-4-dependent macrophage proliferation was observed following parasite infections in mice . Certain miRNAs responding to external stimuli have anti-proliferative or pro-apoptotic effects in immune or cancer cells, which may function as a negative feedback mechanism controlling local cell proliferation. Specifically, E2F1 induces cell cycle progression but also potentiates apoptosis via upregulating pro-apoptotic miR-449a/b expression . Furthermore, Th2-type inflammation induces macrophage proliferation and anti-proliferative miR-378 expression simultaneously, suggesting a complex regulation of in situ macrophage proliferation . miR-342-3p also has pro-apoptotic and anti-proliferative roles in colorectal and breast cancer cells [58, 59, 63]. We found that increased miR-342-3p expression paralleled macrophage proliferation changes observed at the early stage of B. malayi-induced alternative macrophage activation, which suggests a potential role of miR-342-3p in regulating the amount of local viable macrophages . Indeed, our functional studies showed that miR-342-3p over-expression reduced viable macrophage number via induction of apoptosis. We identified a miR-342-3p-repressed anti-apoptotic gene network with well characterized inhibitors of apoptosis, including Bcl2l1, Xiap, Api5, and Birc6 in macrophages by applying a combination of in silico target prediction algorithms and miRNA mimic experiments. In addition, we found that Bcl2l1 is directly repressed by miR-342-3p. These findings strongly suggest a role for IL-4-induced miR-342-3p as a potent negative feedback regulator of macrophage cell number via induction of apoptosis. Thus, the IL-4-triggered proliferative response of macrophages might be accompanied by the simultaneous induction of counteracting cellular processes, generating an endogenous limit of macrophage abundance. The proposed dual role of IL-4 is further supported by the observation that IL-4 is capable of sensitizing macrophages to rapamycin-induced apoptosis . Intriguingly, elevated expression of miR-342-3p in the liver is accompanied by enhanced macrophage apoptosis in malaria-infected mice [93, 94]. These results thus raise the possibility of a connection between macrophage apoptosis and elevated miR-342-3p expression induced by Th2-type inflammation.
The functional plasticity of macrophages determined by polarization signals plays an important role during the development and progression of several human diseases, including sepsis, malignancies, and metabolic disorders . Alternatively activated macrophage-like disease-associated macrophages are observed in the late phase of sepsis and during tumor progression [96, 97]. In addition, an increased number of alternatively activated-like tumor-associated macrophages has been shown to be associated with poor prognosis in several human tumors, including prostate, cervix, breast, and bladder cancer . For these reasons, the transcriptomic and functional characterization of disease-associated macrophages certainly has a high clinical importance. Based on the data presented here and published by others, macrophage-expressed miRNAs show polarization-specific expression patterns and functions [14–17]. The molecular stability of miRNAs in formalin-fixed, paraffin-embedded samples makes them suitable diagnostic and prognostic markers. Furthermore, a well characterized alternative activation-specific marker set has not been available before in the case of human macrophages . Some miRNAs from our study, including miR-342-3p and miR-193b, may be promising candidates as potential biomarkers in combination with other well characterized alternative macrophage activation-specific genes and proteins in human diseases. Recent studies have provided strong evidence that miRNA targeting in vivo using chemically modified oligonucleotides can alter disease outcome in animal models. It would be interesting to examine if in vivo overexpression of pro-apoptotic miR-342-3p has therapeutic relevance in alternative macrophage activation-associated human diseases, including fibrosis and certain malignant tumors that are characterized by pathologic macrophage abundance. However, both diagnostic and therapeutic applications of these miRNAs require further investigation by utilizing disease models and extensive analyses of clinical samples.
The aim of this study was to identify the miRNA signature and its phenotypic consequences in human and mouse models of alternative macrophage activation in vitro and in vivo using a combination of transcriptomic, genomic, bioinformatic, and functional approaches. We uncovered a dynamically regulated miRNA expression pattern during human monocyte–macrophage differentiation and IL-4-mediated alternative macrophage activation. We found that three IL-4-responsive miRNAs (miR-342-3p, miR-125a, and miR-99b) showed conserved regulation in both human and mouse alternatively activated macrophages in vitro and in B. malayi nematode-infected mice in vivo. We also showed that the alternative macrophage activation-specific regulation of these miRNAs is IL-4Rα- and STAT6-dependent. In addition, we determined that both miR-342-3p and its host gene, EVL, are co-regulated directly by STAT6 in mouse and human macrophages. Finally, we demonstrated that macrophage survival was reduced via miR-342-3p-dependent repression of an anti-apoptotic gene network. These results are consistent with the notion that IL-4/STAT6 signaling-induced miR-342-3p is a potent negative feedback regulator of macrophage cell number via induction of apoptosis.
AV, annexin V; BMDM, bone marrow-derived macrophage; ChIP, chromatin immunopreciptiation; FDR, false discovery rate; GO, Gene Ontology; GRO-seq, global run-on sequencing; IL, interleukin; IL-4Rα, IL-4 receptor-α; JAK, Janus kinase; KO, knockout; LPS, lipopolysaccharide; MCSF, macrophage colony stimulating factor; miRNA, microRNA; mRNA, messenger RNA; PI, propidium iodide; PI3K, phosphoinositide 3-kinase; qPCR, quantitative PCR; RMA, robust multi-array average; RT, reverse transcription; STAT1, signal transducer and activator of transcription; TSS, active transcription start site; UTR, untranslated region; WT, wild type.
The authors would like to acknowledge the technical assistance of Ms Ibolya Fürtös, Ms Tímea Cseh, Ms Beáta Szalka, and the comments of Dr Beáta Scholtz and members of the Nagy laboratory on the manuscript. LN is supported by grants from the Hungarian Scientific Research Fund (OTKA K100196 and 116855) and TÁMOP-4.2.2.A-11/1/KONV-2012-0023 VÉDELEM implemented through the New Hungary Development Plan co-financed by the European Social Fund and the European Regional Development Fund. TV is the recipient of a Bolyai Fellowship from the Hungarian Academy of Sciences. CA-G was supported by a grant from the Mexican National Council for Science and Technology (CONACyT grant number 133388). ZC was supported by funding from the European Union, the European Social Fund, and the State of Hungary (TÁMOP 4.2.4. A/2-11-1-2012-0001 ‘National Excellence Program’). DR and JEA were supported by funding from MRC-UK (MR/K01207X/1).
Availability of data and materials
Sequencing data have been submitted to NCBI Sequence Read Archive (SRA) under accession number PRJNA309916. The accession number of GSE71644 is a SuperSeries code for our all Affymetrix miRNA1.0 array data and contains two subsets with following accession numbers: GSE71641, GSE71642. GSE71641 refers to the human peripheral blood monocyte experiment and subseries GSE71642 refers to the RAW264.7 mouse macrophage experiment. ChIP-seq data were downloaded from the NCBI GEO (GSE38379). GRO-seq data were downloaded from the NCBI SRA (SRP019970).
Designed the study: ZC, TV, MK, MP, JEA, ZB, CA-G, and LN. Performed experiments and analyzed data: ZC, TV, MK, COV, QMD-X, DR, ZN, BD, SP, SGT, AH, GN, and EV. The manuscript was written by ZC, MK, and LN with input from the other authors. LN directed the research. All authors provided input to data analysis and approved the final version of the manuscript.
The authors declare that they have no competing interests.
Ethics approval and consent to participate
Animal experiments in the Nagy laboratory were carried out with protocols approved by the Institutional Review Board of the University of Debrecen (file numbers 120/2009/DE MAB and 21/2011/DE MAB). Animal experiments in the Allen laboratory were performed in accordance with the UK Animals (Scientific Procedures) Act of 1986 and approved by the University of Edinburgh Ethical Review Committee. Leukocyte-enriched buffy coats were obtained from healthy blood donors drawn at the Regional Blood Center of the Hungarian National Blood Transfusion Service (Debrecen, Hungary) in accordance with the written approval of the Director of the National Blood Transfusion Service and Institutional Reviewer Board of the University of Debrecen, Faculty of Medicine (OVSzK 3572-2/2015/5200). Written informed consent was obtained from the donors prior to blood donation and their data were processed and stored according to the directives of the European Union.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
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