MIR100 host gene-encoded lncRNAs regulate cell cycle by modulating the interaction between HuR and its target mRNAs

Qinyu Sun; Vidisha Tripathi; Je-Hyun Yoon; Deepak K Singh; Qinyu Hao; Kyung-Won Min; Sylvia Davila; Richard W Zealy; Xiao Ling Li; Maria Polycarpou-Schwarz; Elin Lehrmann; Yongqing Zhang; Kevin G Becker; Susan M Freier; Yuelin Zhu; Sven Diederichs; Supriya G Prasanth; Ashish Lal; Myriam Gorospe; Kannanganattu V Prasanth

doi:10.1093/nar/gky696

. 2018 Aug 8;46(19):10405–10416. doi: 10.1093/nar/gky696

MIR100 host gene-encoded lncRNAs regulate cell cycle by modulating the interaction between HuR and its target mRNAs

Qinyu Sun ^1,², Vidisha Tripathi ^1,^✉,^2,³, Je-Hyun Yoon ^2,^3,^✉, Deepak K Singh ¹, Qinyu Hao ¹, Kyung-Won Min ², Sylvia Davila ², Richard W Zealy ², Xiao Ling Li ⁴, Maria Polycarpou-Schwarz ⁵, Elin Lehrmann ³, Yongqing Zhang ³, Kevin G Becker ³, Susan M Freier ⁶, Yuelin Zhu ⁷, Sven Diederichs ^5,⁸, Supriya G Prasanth ¹, Ashish Lal ⁴, Myriam Gorospe ³, Kannanganattu V Prasanth ^1,^✉

PMCID: PMC6212728 PMID: 30102375

Abstract

Long non-coding RNAs (lncRNAs) regulate vital biological processes, including cell proliferation, differentiation and development. A subclass of lncRNAs is synthesized from microRNA (miRNA) host genes (MIRHGs) due to pre-miRNA processing, and are categorized as miRNA-host gene lncRNAs (lnc-miRHGs). Presently, the cellular function of most lnc-miRHGs is not well understood. We demonstrate a miRNA-independent role for a nuclear-enriched lnc-miRHG in cell cycle progression. MIR100HG produces spliced and stable lncRNAs that display elevated levels during the G1 phase of the cell cycle. Depletion of MIR100HG-encoded lncRNAs in human cells results in aberrant cell cycle progression without altering the levels of miRNA encoded within MIR100HG. Notably, MIR100HG interacts with HuR/ELAVL1 as well as with several HuR-target mRNAs. Further, MIR100HG-depleted cells show reduced interaction between HuR and three of its target mRNAs, indicating that MIR100HG facilitates interaction between HuR and target mRNAs. Our studies have unearthed novel roles played by a MIRHG-encoded lncRNA in regulating RNA binding protein activity, thereby underscoring the importance of determining the function of several hundreds of lnc-miRHGs that are present in human genome.

INTRODUCTION

Human cells utilize only 2% of their genome to generate transcripts with protein-coding sequences. However, a large portion of the genome is transcribed into noncoding RNAs (ncRNAs) with no apparent protein-coding potential. NcRNAs could be broadly classified into two subclasses. Small non-coding RNAs are transcripts smaller than 200 nucleotides, and some well-known examples of small ncRNAs include microRNAs (miRNAs), small interfering RNAs (siRNAs) and Piwi-interacting RNAs (piRNAs). On the other hand, ncRNAs that are larger than 200 nucleotides are defined as long-noncoding RNAs (lncRNAs) (1). Current estimates indicate that human genome harbors >16,000 lncRNA genes (Human GENCODE Release, version 27: http://www.gencodegenes.org/stats/current.html). LncRNA expression is dynamically regulated in a cell-, tissue- or development-specific fashion. Recent studies revealed that lncRNAs play important roles in several biological processes, including cell cycle progression, DNA damage response, stem cell fate determination and X-chromosome inactivation (1–5). In addition, aberrant expression of a large number of lncRNAs is associated with various diseases, including cancer, and a few of the candidate lncRNAs are shown to regulate cancer-related signaling pathways (6–9). At the molecular level, lncRNAs adopt various mechanisms to regulate chromatin organization, gene transcription, and post-transcriptional RNA processing (3). LncRNAs can also serve as molecular scaffolds to modulate nucleic acid-nucleic acid or nucleic acid-protein interactions, or to titrate away proteins and miRNAs from chromatin regions (2,5).

LncRNAs are sub-categorized based on their genomic locations, expression patterns, or functions (10). Some lncRNAs harbor miRNAs within their exonic or intronic sequences, and hence are referred as miRNA-host gene lncRNAs (lnc-miRHGs). miRNAs are short non-coding RNAs (usually 22nt), and they regulate target gene expression post-transcriptionally by promoting mRNA decay or inhibiting translation (11). In the genome, miRNAs are produced from intergenic (28%), intronic (55%), or exonic (17%) regions of host pre-mRNAs or host lncRNAs (12–15). For example, ∼17.5% of miRNAs are produced from lnc-miRHGs (16). The biogenesis and function of miRNAs that are processed from lnc-miRHGs have been well studied. In addition, several lnc-miRHGs show aberrant expression in diseases, hence could serve as important diagnosis or prognosis markers (17–19). However, it is not clear whether the stable and properly spliced pool of lnc-miRHGs, which are processed from the pri-miRHG during miRNA processing plays any vital cellular functions, or merely act as non-functional byproducts of miRNA processing. Very few studies thus far have determined miRNA-independent roles of lnc-miRHGs. For example, PVT1 oncogenic lncRNA, which is processed from the MIRHG harboring miR-1204, miR-1205, miR-1206, miR-1207-5p, miR-1207-3p and miR-1208, positively regulates c-Myc expression and activity (20,21). Similarly, the exon-bearing and completely processed RMST lncRNA (contains miR-1251 within its intronic region) and MIR31HG (contains miR-31 within its intron) are known to play vital roles in neurogenesis and cancer progression, respectively (22,23). Finally, the H19 lncRNA that is processed from a MIRHG plays crucial oncogenic role (19). All of these studies indicate miRNA-independent roles of lnc-miRHGs in various key biological processes.

In the present study, we discovered that multiple lnc-miRHGs, including MIR100HG, are differentially expressed during specific stages of the cell cycle. We observed that the levels of MIR100HG are elevated during G1 stage, and depletion of MIR100HG causes defects in cell cycle progression. More importantly, the spliced, abundant and nuclear-enriched MIR100HG exerts its function in a miRNA-independent manner. We demonstrated that MIR100HG interacts with RNA-binding-proteins (RBPs), such as HuR and several of HuR-target mRNAs. Mechanistic studies indicate that MIR100HG facilitates the interaction between HuR and a subset of its target mRNAs. We conclude that MIR100HG potentially serves as a binding platform for both HuR and its target mRNAs, thus modulating HuR-target mRNA interactions.

MATERIALS AND METHODS

Cell culture

U2OS cells were grown in DMEM containing high glucose, supplemented with Penicillin–Streptomycin (Corning) and 10% fetal bovine serum (FBS) (HyClone, GE). WI-38 was grown in MEM containing high glucose, 10% FBS, and 1% non-essential amino acid (NEA). Cell cycle synchronization of U2OS cells was performed as described previously (24).

Plasmid construction

Full-length MIR100HG (isoform MIR100HG:9 in lncipedia or NR_024430.1 in NCBI) was amplified from U2OS cDNA and was cloned into PGMT-easy vector (Promega) or pCDNA3. Three MIR100HG fragments were sub-cloned from full-length construct into PGMT-easy vector.

Antisense oligonucleotide, 2′MOE and siRNA treatment

Phosphorothioate internucleosidic linkage-modified DNA antisense oligonucleotides (ASOs) were designed and synthesized by Ionis Pharmaceuticals, Inc. They are modified with five 2′-O-methoxyethyl nucleotides on the 5′ and 3′ ends and 10 consecutive oligodeoxynucleotides to support RNase H activity. 2′MOE with phosphorothioate backbone was designed and synthesized by Ionis Pharmaceuticals, Inc., for blocking MIR100HG interaction with mRNAs. ASOs, MOEs and siRNAs (SigmaGenosys, USA) against MIR100HG or HuR were transfected into cells using Lipofectamine RNAiMax (Invitrogen, USA).

cDNA microarray

Total RNA (250 ng) was prepared in triplicate using the RNeasy Mini kit (Qiagen) and labeled using the IlluminaTotalPrep RNA amplification kit (Ambion). Microarrays were performed with the HumanHT-12 v4 Expression BeadChip kit (Illumina). After hybridization, raw data were extracted with Illumina GenomeStudio software, raw probe intensities were converted to expression values using the lumi package in Bioconductor with background correction, variance stabilization and quantile normalization. Differential expression between different conditions was computed by an empirical Bayes analysis of a linear model using the limma package in Bioconductor. Adjusted P-values were calculated with the Benjamini and Hochberg method, and differentially expressed genes were selected with adjusted P-value ≤0.05 and a fold change ≥1.50.

RESULTS

MIR100HG-encoded lncRNAs show elevated levels during G1 phase of the cell cycle

In cancer cells several miRNAs display cell cycle-regulated expression, and are modulated by cell cycle-regulated oncogenic transcription factors including c-Myc (25,26). To determine the involvement of miRNAs and their host genes in cell proliferation, we looked at the expression of several of the c-Myc target MIRHGs, including PVT1, MIR17HG and MIR100HG during the cell cycle (Supplementary Figure S1A). These MIRHG loci or the encoded miRNAs are involved in cell proliferation or cancer progression (27–33). RT-qPCR analyses using primer pairs from exonic-intronic as well as intronic regions in cell cycle synchronized U2OS (osteosarcoma) (Supplementary Figure S1B) cells revealed elevated levels of pri-MIR100HG and PVT1 transcripts, specifically during G1 phase of the cell cycle (Figure 1Aa and Supplementary Figure S1Ca). Pri-MIR17HG RNA showed elevated levels both in the mitotic and G1 stages of the cell cycle (Supplementary Figure S1Cb). Elevated PVT1 expression is highly correlated with cell proliferation and tumor formation (21). MIR17-92 miRNA cluster is a known oncomir (34,35), and MIR100HG-encoded miRNAs (miR-100, let7a2 and miR-125b1) are known to regulate cell proliferation (27–33). These results indicate that the expression of these lnc-miRHGs is dynamically regulated during the cell cycle.

MiR-100, let7a2 and miR-125b1 are embedded within the last intron of MIR100HG (Figure 1B and Supplementary Figure S1Ac, Figure 1B showing isoform MIR100HG:9 in lncipedia or NR_024430.1 in NCBI). It is interesting to note that MIR100HG-encoded miRNAs are shown to play pro- as well as anti-proliferative roles. For example, let-7 miRNA family is a known tumor suppresser miRNA, and acts as a negative regulator of cell growth. Both miR-125-b1 and miR-100 act as oncogenic or tumor suppressor miRNAs in various cancers (27–33). Because of this, we were interested in determining whether the exon-bearing lncRNA/s transcribed from MIR100HG locus play any role in cell proliferation. MIR100HG produces several isoforms of MIR100HG lncRNAs (in the rest of the manuscript, we refer MIR100HG for exon-bearing lncRNAs from MIR100HG locus) due to alternative splicing and differential promoter usage (Supplementary Figure S1Ac). In order to test the relative levels of the spliced MIR100HG during the cell cycle, we performed RT-qPCR analyses in RNA samples from synchronized U2OS cells using exon–exon junction primer pairs to amplify the exonic region (exon 2–exon 3 and exon 3–exon 4) shared by most of the MIR100HG isoforms. Similar to what we observed in the case of pri-miR100HG RNA, even the spliced MIR100HG showed elevated levels, specifically during G1 (Figure 1Ab and Ac). Exonic primers targeting the last exon (both spliced and pri-miR100HG transcripts) also displayed same expression pattern (Supplementary Figure S1Cc). MIR100HG was found to be a predominantly nuclear and relatively stable lncRNA with a half-life of ∼75 min (Figure 1C, Supplementary Figure S1Da). Interestingly, MIR100HG stability was regulated during cell cycle, as observed by its enhanced stability during G1 phase over S-phase of the cell cycle (Supplementary Figure S1Db).

Next, we tested the levels of mature and precursor miR-100 and miR-125b1 (both are generated from MIR100HG locus) during the cell cycle by using Taqman and cleavage assay (an assay that quantifies the production of pre-miRNA) respectively (36). The levels of mature, pre-miRNA and pri-miRNAs did not change significantly during the cell cycle (Figure 1Da and Db, Supplementary Figure S1E). The absence of much fluctuation of mature miRNAs during the cell cycle could be attributed to their enhanced stability. The dynamic expression and differential stability of the nuclear-restricted MIR100HG during the cell cycle implies that the lncRNAs produced from the MIR100HG locus might participate in cell cycle progression.

Elevated levels of MIR100HG were previously observed in megakaryoblastic leukemia (AMKL) cell lines, and high levels of MIR100HG were correlated with poor prognosis in cases of cervical cancer patients (37,38). We therefore determined the levels of MIR100HG in various cancer cell lines and in non-tumorigenic cells. We compared the level of MIR100HG among human diploid lung fibroblast (WI-38), cervical cancer (HeLa), bone osteosarcoma (U2OS) and highly tumorigenic and metastatic MCF10CA1a.cl1 breast cancer (M4) cell lines (39,40). Compared to the three cancer cell lines, non-tumorigenic diploid WI-38 cells showed lower levels of MIR100HG (Supplementary Figure S1Fa). In addition, we also determined the levels of MIR100HG in a MCF10A-derived isogenic breast cancer progression cell line model system, represented by M1-M4 cell lines. M1 represents non-tumorigenic MCF10A cells, and M2 represents HRAS-expressing M1 cells but show very low tumorigenic potential when implanted in immune compromised mice. M3 and M4 were isolated from tumor samples from mice that were xenografted with M2 cells. M3 cells are highly tumorigenic and form well-differentiated tumors in mice, but they display low metastatic potential. On the other hand, M4 cells form highly undifferentiated tumors and very efficiently metastasize in the lungs (39–43). We observed that MIR100HG levels were dramatically increased in both M3 and M4 cells compared to M1 and M2 cells (Supplementary Figure S1Fb). These results suggest that G1-phase upregulated MIR100HG tends to show elevated expression in cancer cells. Publicly available RNA-seq data sets from U2OS cells showed expression of MIR100HG (Supplementary Figure S1G). Further, RNA copy number analyses revealed that asynchronous U2OS and M4 cells contained ∼68 and ∼365 copies of processed MIR100HG lncRNAs respectively (Supplementary Figure S1H). Northern blot (NB) using a probe from exon 4 of MIR100HG showed a specific signal of ∼3.5 kb (Figure 1Ea). NB using a probe comprising of exons 1–3 showed two discrete bands, one at 3.5 kb and another weak band at 5 kb (Figure 1Eb), indicating that MIR100HG produces multiple isoforms of MIR100HG via alternative splicing (please also see Supplementary Figure S1Ac). Finally, we determined the protein-coding potential of MIR100HG isoforms utilizing the integrated database of annotated human lncRNAs: Lncipedia.org (https://lncipedia.org/), and identified MIR100HG as a non-coding transcript (Supplementary Figure S1I). In summary, we identified a stable, abundant, spliced, G1 phase-enriched, and nuclear restricted lncRNA, transcribed from the MIG100HG locus that showed elevated expression in several of the tested cancer cell lines.

MIR100HG regulates cell cycle progression

To determine the involvement of MIR100HG during the cell cycle, we successfully depleted MIR100HG by transfecting U2OS cells with modified DNA antisense oligonucleotides (ASO1 and ASO2) (Figure 2A and Supplementary Figure S2A) for 48 h. MIR100HG-depleted cells did not show any significant change in the total cellular levels of miR-100, pre and pri-miR-100, and pre-miR-125b1 post 48 h of ASO transfection (Figure 2B, Supplementary Figure S2B). In addition, we examined the RNA levels of several known miR-100 target mRNAs in control and MIR100HG-depleted cells (31,44–47). RT-qPCR and immunoblot analyses results revealed that both control and MIR100HG-depleted cells showed comparable levels of several of the tested miR-100 target mRNAs (Figure 2C and Supplementary Figure S2C). These results confirm that cells depleted of MIR100HG for 48 h did not show changes in either the levels or activity of the MIR100HG intron-encoded miRNAs. It is possible that miR-100 and miR-125b1 are stable miRNAs, therefore the depletion of MIR100HG for 48 h did not affect the steady-state levels of these miRNAs. Alternatively, MIR100HG gene locus could produce different sets of transcripts, some of which will be processed to synthesize exon-bearing lncRNAs, and the others act as the microRNA host transcripts.

Figure 2. — *MIR100HG* depletion results in cell cycle arrest. (A) RT-qPCR to determine the relative levels of *MIR100HG* in U2OS cells transfected with control (SCR) or antisense oligonucleotides targeting *MIR100HG* (ASO1, ASO2). (B) Taqman RT-qPCR assay to reveal relative levels of miR-100 in control (SCR) and *MIR100HG*-depleted (ASO1 and ASO2) cells. Right: RNA cleavage assay to determine the levels of pre-miR-100 and pre-miR-125b1 in control and *MIR100HG*-depleted cells. (C) RT-qPCR to determine the relative levels of several miR-100 target gene mRNAs in control (SCR) and *MIR100HG*-depleted (ASO1 and ASO2) cells. (D) Flow cytometry analyses in control (SCR) and *MIR100HG*-depleted (ASO1 and ASO2) U2OS cells. Quantification was performed using FCS Express. *P≤ 0.05, **P≤ 0.01, ***P ≤ 0.001, ****P ≤ 0.0001 by two-tailed Student's t-test, n = 3. Error bars represent standard deviation.

To determine the involvement of MIR100HG in cell proliferation, we performed flow cytometry analysis and found that MIR100HG-depleted U2OS cells showed elevated levels of G2/M population with a concomitant reduction in G1 population (ASO1 and ASO2, Figure 2D). In addition, depletion of MIR100HG in U2OS cells using a third ASO from exon 2 (ASO3; Supplementary Figure S2A) also showed a similar phenotype (Supplementary Figures S2D and S2E). We also depleted MIR100HG using an independent siRNA targeting exon 4 (Supplementary Figures S2A and S2F). Cells depleted of MIR100HG using siRNA also showed G2/M arrest without affecting the mature miR-100 levels, confirming the specificity of the cell cycle phenotype (Supplementary Figures S2G and S2H). Finally, we performed rescue experiments by overexpressing the full-length spliced MIR100HG isoform in cells depleted of the endogenous MIR100HG. We observed that the MIR100HG cDNA could partially rescue the G2/M arrest phenotype (Supplementary Figure S2I). This data confirmed the specificity of the knockdown phenotype, and further indicated microRNA-independent cell cycle role for lncRNAs processed from the MIR100HG locus.

In addition to G2/M arrest, MIR100HG depletion in diploid fibroblasts (WI-38) also resulted in defective S phase progression, as observed by reduced bromo-deoxy uridine (BrdU) uptake in the BrdU/propidium iodide (PI) flow cytometry analysis (Supplementary Figure S2J). In order to determine the rate at which cells progress through S phase, we synchronized control and MIR100HG-depleted cells in G1/S using aphidicolin, and released them into S phase for the indicated time points (4, 8 and 12 h post-aphidicolin release). We found that, when compared to control ASO-treated cells, MIR100HG-depleted cells displayed a delay in their progression through S phase (Supplementary Figure S2K). This was obvious in the 8 h release time point in which the control cells had progressed into G2/M phase, but a fraction of MIR100HG-depleted cells continued to have DNA content between 2C and 4C (observed by the broader peak), implying the slow S-phase progression. Similarly, a proportion of control cells progressed into G1 phase at the 12 h release time point. However, most of the MIR100HG-depleted cells at the 12 h time point accumulated in G2/M phase and failed to enter G1 phase.

In order to gain insight into the gene expression program that is affected in MIR100HG-depleted cells, we performed transcriptome microarray in control and MIR100HG-depleted U2OS cells. We identified 722 downregulated genes (≤0.67 fold) and another 577 upregulated genes (≥1.5-fold) in MIR100HG-depleted cells. Pathway analyses revealed that these genes, whose expression levels are altered in MIR100HG-depleted cells, play vital roles in crucial pathways controlling cell growth, including cell cycle, cell proliferation, cell death and survival, further supporting the cellular phenotype observed in MIR100HG-depleted cells (Supplementary Table S1, Supplementary Figure S2L).

In summary, we found that human diploid and cancer cells depleted of MIR100HG showed defects in cell cycle progression, as observed by slow S phase progression and/or G2/M arrest. We propose that that G1-upregulated MIR100HG might play a crucial role in the entry of cells into G1 phase of the cell cycle, and its depletion results in defects in G1 entry, resulting in cells being arrested in G2/M phase of the cell cycle.

MIR100HG interacts with HuR and influences the association of HuR with its target mRNAs

It is known that several nuclear-retained lncRNAs regulate gene expression by influencing the localization and/or activity of various RBPs (5,48–50). In order to gain mechanistic insights into the role of MIR100HG, we performed RNA-affinity pull-down in U2OS nuclear extracts using biotin-labeled MIR100HG followed by mass-spectrometry (Figure 3A). Mass spectrometry showed potential interaction between MIR100HG and several RNA binding proteins (RBPs) (Supplementary Figure S3A). Further, we validated the interaction between MIR100HG and RBPs such as HuR, PTB and PUF60, by biotin–RNA pull-down followed by immunoblotting using nuclear extracts (Figure 3B). HuR, PTB, PUF60, but not AUF1, showed positive interaction with MIR100HG.

Figure 3. — *MIR100HG* interacts with RNA binding protein HuR. (A) Representative silver-stained SDS-PAGE gel showing specific bands of the extracts incubated with biotin-labeled *MIR100HG* followed by RNA pull-down assay. Nuclear extract of U2OS was used in biotin–RNA pulldown assay. RNA Pull-down using biotin-labeled YFP RNA (lane 2, YFP) or beads only without RNA (lane 3, Beads) served as controls. (B) *MIR100HG*-interacting proteins identified by mass spectrometry analyses were validated by *in vitro* biotin–RNA pull-down followed by immunoblotting. RNA Pull-down was performed using biotin-labeled full-length *MIR100HG*. YFP and beads serve as control. (C) HuR-RIP or IgG-RIP followed by RT-qPCR during specific stages of cell cycle (G1, G1/S, S, G2, M phases) to determine the interaction between HuR and *MIR100HG*. RT-qPCR data was normalized over GAPDH mRNA. (D) Schematic showing three fragments of *MIR100HG*. (E) *In vitro* biotin–RNA pull-down followed by immunoblotting to detect the interaction between HuR and specific fragments (1, 2 and 3) or full length (FL) *MIR100HG*. YFP RNA was used as negative control. SRSF1 served as a negative control. *P≤ 0.05, **P≤ 0.01, ***P ≤ 0.001, ****P ≤ 0.0001 by two-tailed Student's t-test, n = 3. Error bars represent standard deviation.

HuR is a ubiquitous member of neuronal ELAV-like proteins. It recognizes AU-rich elements (ARE) or U-rich sequences, typically within the 3′-untranslated region (UTR) of RNAs (51–53). HuR is known to control multiple biological events including the cell cycle, apoptosis, and cancer progression (54–58). We therefore investigated the functional significance of the interaction between HuR and MIR100HG. We first confirmed the interaction between HuR and endogenous MIR100HG by HuR ribonucleoprotein (RNP) immunoprecipitation (RIP) analysis in cell cycle synchronized cell extracts followed by RT-qPCR. HuR showed enhanced interaction with MIR100HG during the G1/S phase of the cell cycle (Figure 3C). Immunoblotting in cell cycle synchronized cell extracts indicated that the level of HuR in G1/S was comparable to most other cell cycle stages except G1 phase, where the levels were found to be relatively low (Supplementary Figure S3B). In order to map the sequence elements within MIR100HG that interact with HuR, we performed biotin–RNA pull-down followed by Immunoblotting using 3 partially overlapping fragments of MIR100HG (Figure 3D). We observed that both fragments 2 and 3 within MIR100HG interacted with HuR (Figure 3E). Both the HuR-interacting fragments were from the 3′end of the last exon (exon 4) of MIR100HG. On the other hand, another RBP SRSF1 failed to interact with any of the fragments, confirming the specificity of HuR: MIR100HG interaction (Figure 3E). Interestingly, we found that fragments 2 and 3 contain two U-rich repeats (18 and 15 nt long U-rich repeats respectively, Figure 3D). Based on these data, we conclude that HuR preferentially interacts with the last exon of MIR100HG.

HuR is known to regulate the stability or translation of many mRNAs (51,53). To determine whether MIR100HG influences the association of HuR with its target mRNAs, we performed HuR RIP followed by microarray using RNA from control and MIR100HG-depleted U2OS cells (Supplementary Figure S4A). Interestingly, knockdown of MIR100HG significantly decreased the association of HuR with several of its target mRNAs (Supplementary Figures S4B and S4Ca, Supplementary Table S2). Further, by performing HuR-RIP followed by RT-qPCR in control and MIR100HG-depleted cells, we confirmed the reduced association of HuR with several of its target mRNAs (MSH3, CCR6, and CCND2; Figure 4A). At the same time, MIR100HG depletion did not alter the association of HuR to several constitutively expressed housekeeping gene mRNAs (Supplementary Figure S4Cb). Our results imply that MIR100HG positively regulates HuR association with several of its target mRNAs.

Figure 4. — *MIR100HG* regulates the interaction between HuR and its target mRNAs. (A) HuR RIP followed by RT-qPCR in control (SCR) and *MIR100HG*-depleted (ASO1) cells to determine the interaction between HuR and targets, including (a) MSH3 mRNA, (b) CCR6 mRNA and (c) CCND2 mRNA. RT-qPCR data was normalized to GAPDH mRNA. (B) RT-qPCR to determine the relative levels of several of the HuR-target mRNAs in control (SCR) and *MIR100HG*-depleted (ASO1 & ASO2) U2OS cells. (C) RT-qPCR to determine the relative levels of *MIR100HG* in control and HuR-depleted cells. (D) Immunoblot to detect the relative levels of HuR and protein levels of its target mRNAs in control (SCR) and *MIR100HG*-depleted (ASO1) cells (a), and in control (Ctr-si) and HuR-depleted (HuRsi) cells (b). α-Tubulin was included as loading control. The relative protein amount in knockdown samples (with 100% loading) was normalized relative to α-Tubulin, and compared to control. Quantification was performed using ImageJ, quantification results for three repeats is shown in Supplementary Figure S4H. (E) Immunoblot to detect the relative protein level of CCND2 in U2OS cells transfected with empty vector or *MIR100HG* overexpression vector (same plasmid as in Supplementary Figure S2I), with control or *MIR100HG* antisense oligo (ASO1). Quantification was performed using ImageJ. α-Tubulin was included as loading control. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001, ****P ≤ 0.0001 by two-tailed student's t-test. For (Aa-b, B, E), n = 3. For (Ac), n = 2. Error bars represent standard deviation.

It is possible that the aberrant association of HuR to its target mRNAs in MIR100HG-depleted cells could be due to reduced levels of these mRNAs. To test this, we checked the steady-state levels of HuR target mRNAs in control and MIR100HG-depleted cells using our microarray data set. Depletion of MIR100HG did not affect steady-state levels of known HuR target mRNAs (Supplementary Table S1). Microarray results were further validated by RT-qPCR analyses (Figure 4B). We also found that knockdown of MIR100HG did not alter either the total levels or the subcellular distribution of HuR (Supplementary Figures S4D and S4E). These findings indicated that the reduction of HuR’s association with a subset of its target mRNAs detected upon MIR100HG depletion was not due to changes in total levels or localization of HuR, but due to changes in its ability to interact with specific mRNA targets.

It is interesting to note that the strongest interaction between MIR100HG and HuR was observed at the G1/S-boundary (Figure 3C). MIR100HG level was also relatively low during the G1/S transition (Figure 1A). Although HuR is known to stabilize mRNAs in many instances, recent studies from several laboratories, including ours, have shown that HuR can also destabilize mRNAs and lncRNAs (59–61). In order to determine whether HuR influences the levels of MIR100HG, we compared the levels of MIR100HG in control and HuR-depleted cells. HuR-depleted cells showed a small but significant increase in the levels of MIR100HG, indicating that the association of HuR with MIR100HG at the end of G1 phase could also facilitate the destabilization of lncRNA (Figure 4C).

HuR participates in several crucial post-transcriptional processes such as mRNA stability, and is also known to regulate translation (62). We tested whether MIR100HG- or HuR-depleted cells showed any change in the mRNA stability or protein levels of a few HuR targets. Both MIR100HG and HuR-depleted cells did not show any change in the stability of the tested mRNAs (Supplementary Figures S4F and S4G). Interestingly, MIR100HG- or HuR-depleted cells showed decrease in the protein levels of CCND2/Cyclin D2, a known HuR target (Figure 4Da and Db, Supplementary Figure S4H). Furthermore, we rescued the levels of CCND2 in MIR100HG-depleted cells by exogenously expressing one of the MIR100HG isoforms (Figures 1B and 4E). MIR100HG seems to regulate the cellular levels of CCND2 protein. It is possible that reduced interaction between HuR and CCND2 mRNA observed in MIR100HG-depleted cells contributes to reduced levels of CCND2 protein.

U-rich domains within MIR100HG modulate the interaction between MIR100HG and HuR RNP complex

Next, we probed into the potential mechanism utilized by MIR100HG to influence the interactions between HuR and its target mRNAs. LncRNAs is known to function as scaffold to stabilize RNA: protein, DNA: protein, and protein: protein interactions (2). We determined whether MIR100HG, by interacting with both HuR and its target mRNAs, modulates HuR-target mRNA association or stabilizes HuR-target mRNA RNP complex assembly. To test this, we used computational prediction tools (63) to find mRNAs that have sequence complementarity with MIR100HG. From the prediction analysis, we narrowed down the candidate mRNAs containing at least 18nt of sequence complementarity with MIR100HG with >88% identity. From the prediction, we found that 7706 mRNAs met these criteria (Supplementary Table S3), while only 24 of them were identified as HuR-interacting mRNAs by our HuR RIP analysis (Supplementary Table S3). Our results suggested that HuR and MIR100HG could bind to several shared target mRNAs. Interestingly, most predicted interactions between mRNAs and MIR100HG occurred through the first U-rich repeat sequences that present in the last exon of MIR100HG (Figure 5A). The low sequence complexity of 18-nt U-repeat indicated low specificity. However, due to the fact that MIR100HG interacted with HuR, we asked if MIR100HG displayed higher binding affinity to HuR target mRNAs among all of these predicted mRNAs. To test this possibility experimentally, we incubated biotin-labeled MIR100HG or YFP mRNA (as a negative control) in total cellular extracts, and performed in vitro pull-down followed by RT-qPCR to identify RNAs that interact with MIR100HG. We observed significant interaction between MIR100HG and either MSH3, CCR6, or CCND2 mRNAs, compared to biotin-labeled YFP RNA. (Figure 5B, MIR100HG+MOE-SCR). These three mRNAs were previously shown to interact with HuR in a MIR100HG-dependent manner (Figure 4A). Absence of specific association between MIR100HG and GPI mRNA confirmed the specificity of the interaction (Supplementary Figure S5A).

Figure 5. — U-rich domains within *MIR100HG* facilitates the interaction between *MIR100HG* and HuR or HuR-target mRNAs. (A) Schematic showing the position of two U-rich regions and the 2′-O-methoxyethyl antisense oligonucleotide (MOE, A-rich) within the last exon of *MIR100HG*. (B) *In vitro* biotin–RNA pull-down followed by RT-qPCR to determine the interaction between *MIR100HG* and HuR-target mRNAs in SCR and U-rich targeting MOE-treated extracts. RT-qPCR data was normalized to GAPDH mRNA. Relative fold enrichment of (a) MSH3 mRNA, (b) CCR6 mRNA, and (c) CCND2 mRNA were calculated by comparing over biotin-YFP, which was used as negative control. A non-*MIR100HG*-interacting example GPI mRNA is shown in Supplementary Figure S5A. (C) *In vitro* biotin–RNA pull-down followed by immunoblotting to detect the association between *MIR100HG* and HuR in control (MOE-SCR) and *MIR100HG* targeting (MOE-100) MOE pre-incubated samples. YFP RNA was used as negative control. SRSF1 served as a negative protein control. biotin–RNA was pre-incubated with 2′-MOE oligos (MOE-SCR & MOE-100) prior to *in vitro* pull-down. (D) HuR RIP followed by RT-qPCR in in SCR and U-rich targeting MOE (MOE-100) treated cells to determine the interaction between HuR and targets, including (a) MSH3 mRNA and (b) CCND2 mRNA. RT-qPCR data was normalized to GAPDH mRNA. Fold of enrichment (relative to IgG) is plotted. (E) Immunoblot to detect the protein levels of HuR target genes in control (SCR) and MOE-100 transfected cells. α-Tubulin was used as loading control. The relative fold of protein amount in MOE-100-treated samples (with 100% loading) was normalized over α-Tubulin, and compared to MOE-SCR. Quantification was performed using Image J and quantification results for three repeats is presented in Supplementary Figure S5B. (F) Model depicting the potential mode of action of *MIR100HG. MIR100HG* lncRNA interacts with both HuR and HuR target mRNAs and increases their local concentration, and thus facilitating HuR-target mRNA interactions. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001, ****P ≤ 0.0001 by two-tailed Student's t-test, n = 4 for 5B, n = 3 for all other figures. Error bars represent standard deviation.

We next asked if the U-repeat region within the MIR100HG is essential for facilitating the lncRNA: mRNA interactions. To answer this question, we designed a 20 nt-long 2′-O-methoxyethyl antisense oligos (MOE; MOE-100), which is complementary to the U-rich repeat region within the MIR100HG (Figure 5A). For the assay, we pre-incubated biotin-MIR100HG with either control or this A-rich MOE (MOE-100); then incubated the RNA with cellular extract, and finally determined the efficiency with which MIR100HG interacted with MSH3, CCR6, CCND2 or GPI mRNAs. Pre-incubation of biotin-labeled MIR100HG with MOE, which hybridized to U-rich sequences within the lncRNA, completely abolished the interaction between MIR100HG and either MSH3, CCR6 or CCND2 mRNAs (Figure 5B and Supplementary Figure S5A, MIR100HG+MOE-100). These results reveal that the U-rich sequences that present in the 3′ end of MIR100HG promotes its interaction with specific members of HuR-target mRNAs.

We also determined whether the U-rich sequence element also facilitates the interaction between MIR100HG and HuR. For this, we performed RNA-affinity pull-down in cell extracts using biotin-labeled MIR100HG which was pre-incubated with biotin-labeled control (SCR) or A-rich MOE (MOE-100) oligos followed by immunoblotting. HuR showed strong interaction with MIR100HG only in SCR-MOE pre-incubated extracts and not in A-rich MOE-pre-incubated extracts (Figure 5C), indicating that MIR100HG utilizes the U-rich motif to interact with both HuR and HuR-target mRNAs. Finally, we were interested to see the involvement of MIR100HG U-rich sequence element in the in vivo interaction between HuR and its target mRNAs. Towards this, we performed HuR RIP followed by RT-qPCR in cells transfected with control and A-rich MOE-oligos to determine the interaction between HuR and its target mRNAs. We found that the A-rich MOE (MOE-100), which could disrupt the interaction between MIR100HG and HuR and HuR target mRNAs, also attenuated the interaction between HuR and its target mRNAs, including MSH3 and CCND2 (Figure 5D). Finally, we also found that the protein levels of CCND2 and MSH3 were reduced in cells transfected with the A-rich MOE-oligos. (Figure 5E, Supplementary Figure S5B). These results suggest that the U-rich sequences within MIR100HG are the necessary for MIR100HG’s regulatory role in HuR activity modulation. Our results reveal that the interaction between MIR100HG and HuR or its target mRNAs modulates HuR-association to its target mRNAs.

DISCUSSION

Human lncRNAs play crucial roles in several cellular processes such as cell proliferation and differentiation. Most lnc-miRHG studies have focused on understanding the mechanisms whereby cells synthesize miRNAs from their host transcripts. Very few studies thus far have looked at the potential miRNA-independent roles and the mechanism of action of lncRNAs that are processed from the exonic regions of MIRHG primary transcripts. In the present study, we provided evidence for the miRNA-independent role for a lnc-miRHG. We found MIR100HG levels to be elevated during G1 phase of the cell cycle, and cells depleted of MIR100HG showed defects in their entry into G1, supporting MIR100HG’s role in cell cycle progression. Furthermore, mechanistic studies revealed that MIR100HG acts as a scaffold to modulate the interaction between HuR and its target mRNAs.

It is generally believed that the role of MIRHG is to produce miRNAs, and the lncRNAs that are processed from MIRHG primary transcripts do not execute any significant role. However, few recent studies have reported miRNA-independent roles played by lnc-miRHGs. (20,22–23,36). These studies underscore the functional significance of this class of lncRNAs, which is ignored before, and indicate the importance of understanding the independent roles played by MIRHG-encoded lncRNAs. Our study demonstrates the involvement of MIR100HG in the cell cycle and its role in regulating the affinity of an RBP to its target mRNAs, which further underscores the importance of determining the role of several hundreds of lnc-miRHGs in human genome.

RNA pull-down followed by mass spectrometry analyses revealed that MIR100HG interacts with several RBPs, including HuR, PTB and PUF60. HuR is a ubiquitously expressed member of the Hu/ELAV family of proteins. It was initially described as an adaptor protein that influences mRNA export (64). High-throughput protein-RNA interaction studies identified several hundreds of RNAs that interact with HuR (52,53). HuR is found to regulate several vital processes such as mRNA stability, translation, and miRNA biogenesis (62,64–65). Besides MIR100HG, HuR is shown to interact with several other lncRNAs, including lincRNA-p21, UFC1, OIP5-AS1 and linc-MD1, and influence their activity (36,61,66–67). For example, HuR binding to linc-MD1 prevents Drosha-mediated cleavage and production of pre-miRNA from the linc-MD1 transcript (36). The 3′ end of MIR100HG contains two independent U-rich sequence elements, and our in vitro pull-down assays demonstrated that HuR interacts with the U-rich sequence elements. Interestingly, MIR100HG also utilizes the same U-rich sequence elements to interact with several tested HuR target mRNAs. It is possible that MIR100HG could simultaneously interact with both HuR as well as its target mRNA through the U-rich elements. By this, MIR100HG bring both HuR and its target mRNA in close proximity, and this can further increase HuR and target mRNA interactions. The reduced interaction between HuR with target mRNAs in MIR100HG-depleted cells further strengthens our hypothesis.

Lnc-miRHGs are generally very unstable in nature (16), which indicates that these lncRNAs are degraded rapidly after generating miRNAs, and thus they might not serve other functions. On the other hand, MIR100HG was found to be an abundant and stable lncRNA. RNA stability assays revealed that MIR100HG showed enhanced stability during G1 phase of the cell cycle. Interestingly, MIR100HG showed increased association with HuR during the time window that also coincides with its decreased stability. Furthermore, HuR-depleted cells showed an increase in the level of MIR100HG, indicating that HuR destabilizes MIR100HG. In support of this argument, recent studies from several laboratories, including ours, have provided evidence demonstrating the role of HuR as an RNA-destabilizing factor (60–61,68–70). For example, HuR is known to promote the degradation of lincRNA-p21 by facilitating the recruitment of let-7b/AGO2 complex to the lncRNA (61).

Loss-of-function studies indicate that MIR100HG is required for cell cycle progression. Our results are consistent with a previous study, which reported that MIR100HG positively regulates cell proliferation in leukemia cells (37). Interestingly, we observed that the levels of only the MIR100HG, and not the intron-embedded miRNAs (miR-100 and miR-125b1), were elevated during G1 phase of the cell cycle. Since depletion of MIR100HG did not alter the levels of either the MIRHG-encoded miRNAs or the mRNAs that are targets of these miRNAs, we concluded that lncRNA MIR100HG plays a miRNA-independent role in regulating cell cycle progression. In addition, MIR100HG rescue experiments reveal that the spliced MIR100HG lncRNA plays a crucial role in cell cycle progression. The MIR100HG gene locus synthesizes multiple MIR100HG lncRNA transcripts by utilizing multiple promoters. It is possible that only a subset of the isoforms is processed to produce microRNAs whereas rest of them essentially function as host gene of the spliced lncRNAs. Presently, it is not clear how MIR100HG regulates cell cycle progression. One possibility could be that MIR100HG promotes the interaction between HuR and mRNAs of cell cycle genes. Such association is crucial for the proper functioning of those mRNAs. HuR is known to regulate cell division and checkpoint response through multiple mechanisms, including modulating the stability and translation of key cell cycle regulators, such as cyclins (58). The localization of HuR is also regulated during the cell cycle (71,72), and such dynamic cellular distribution could influence its interaction with target mRNAs as well as other lncRNAs such as MIR100HG. To test the involvement of MIR100HG in modulating HuR activity, we determined the mRNA stability and protein levels of HuR targets such as MSH3 and CCND2 mRNAs in cells depleted of MIR100HG. HuR-depleted cells were used as controls in these experiments. To our surprise, we did not see any significant change in the stability of both these transcripts in cells depleted of either MIR100HG or HuR, indicating that HuR does not seem to regulate the stability of these mRNAs. Immunoblot analyses revealed a consistent decrease in the levels of CCND2, and not MSH3 protein, in MIR100HG or HuR-depleted cells. This suggests that MIR100HG does not modulate the outcome of all of the HuR-target mRNAs in the similar fashion.

Since HuR interacts with several hundred mRNAs (52), and MIR100HG-depleted cells showed defects in the association between HuR and a large number of its targets, it would be challenging to identify a single specific target mRNA whose aberrant function contributes to the observed cell cycle arrest upon MIR100HG depletion. Alternatively, MIR100HG regulates the cell cycle via modulating the activity of other RBPs, and future efforts will be made to test this possibility. In summary, we demonstrated that MIR100HG regulates cell cycle progression. Furthermore, we observed that MIR100HG interacted with HuR and its target mRNAs in a cell cycle-regulated manner. The U-rich regions within the MIR100HG were found to be essential for its interaction with HuR as well as with HuR target mRNAs. Based on this, we propose a model (Figure 5F), whereby the abundant MIR100HG interacts with both HuR and its target mRNAs through its U-rich domains. Such interaction further facilitates the association between HuR and its target mRNAs. By this mechanism, MIR100HG acts as a ‘nucleating site’ or ‘scaffold’ to increase the local concentration of HuR and its target mRNA, thus facilitating HuR target mRNA interactions. In summary, our studies have unearthed novel roles played by a MIRHG-encoded lncRNA in regulating RBP activity, thereby underscoring the importance of determining the function of several hundreds of lnc-miRHGs that are present in the human genome.

DATA AVAILABILITY

Sequence data of both cDNA microarray and HUR-RIP microarray have been submitted to NCBI Gene Expression Omnibus (GEO) (http://www.ncbi.nlm.nih.gov/geo/). The accession numbers are: GSE108280 (cDNA microarray) and GSE107836 (HuR RIP).

Supplementary Material

Supplementary Data

Click here for additional data file.^{(10.4MB, zip)}

ACKNOWLEDGEMENTS

We would like to thank K.V. Prasanth and S.G. Prasanth lab members for their comments and suggestions. We thank Drs Eric Bolton (UIUC) and M. Hastings (Rosalind Franklin University) for sharing reagents, and Dr P. Yao for mass spectrometry analyses. We thank J. Roy and S. Sudhakar for proof reading the manuscript.

SUPPLEMENTARY DATA

Supplementary Data are available at NAR Online.

FUNDING

National Institute of Health [1RO1GM088252 to K.V.P., 1RO1GM099669 to S.G.P.]; American Cancer Society [RSG-11-174-01-RMC to K.V.P.]; National Science Foundation [career, 1243372 to S.G.P. and EAGER 1723008 to K.V.P.]; National Institute on Aging intramural Research Program, NIH [Z01-AG000511 to M.G.]; Medical University of South Carolina and Hollings Cancer Center (to J.H.Y.); National Institute on Cancer Intramural Research Program, NIH (to A.L.). The funders had no role in study design, data collection and analysis. Funding for open access charge: Start-up grant from UIUC.

Conflict of interest statement. S.M.F. is an employee of Ionis Pharmaceuticals, and receives salary from the company.

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PERMALINK

MIR100 host gene-encoded lncRNAs regulate cell cycle by modulating the interaction between HuR and its target mRNAs

Qinyu Sun

Vidisha Tripathi

Je-Hyun Yoon

Deepak K Singh

Qinyu Hao

Kyung-Won Min

Sylvia Davila

Richard W Zealy

Xiao Ling Li

Maria Polycarpou-Schwarz

Elin Lehrmann

Yongqing Zhang

Kevin G Becker

Susan M Freier

Yuelin Zhu

Sven Diederichs

Supriya G Prasanth

Ashish Lal

Myriam Gorospe

Kannanganattu V Prasanth

Abstract

INTRODUCTION

MATERIALS AND METHODS

Cell culture

Plasmid construction

Antisense oligonucleotide, 2′MOE and siRNA treatment

cDNA microarray

RESULTS

MIR100HG-encoded lncRNAs show elevated levels during G1 phase of the cell cycle

Figure 1.

MIR100HG regulates cell cycle progression

Figure 2.

MIR100HG interacts with HuR and influences the association of HuR with its target mRNAs

Figure 3.

Figure 4.

U-rich domains within MIR100HG modulate the interaction between MIR100HG and HuR RNP complex

Figure 5.

DISCUSSION

DATA AVAILABILITY

Supplementary Material

ACKNOWLEDGEMENTS

SUPPLEMENTARY DATA

FUNDING

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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