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miR-130a alleviates neuronal apoptosis and changes in expression of Bcl-2/Bax and caspase-3 in cerebral infarction rats through PTEN/PI3K/Akt signaling pathway
research-article
Author(s):
Yinming Wang ,
Junquan Gu ,
Linlin Hu ,
Liang Kong ,
Tinggang Wang ,
Meiqi Di ,
Chaosheng Li ,
Shuhua Gui
Publication date (Electronic):
03 January 2020
Publisher:
D.A. Spandidos
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Abstract
Effect of micro ribonucleic acid (miR)-130a on neuronal apoptosis in rats with cerebral infarction (CI) was studied to explore whether phosphatase and tensin homolog deleted on chromosome ten (PTEN)/phosphatidylinositol 3-hydroxy kinase (PI3K)/protein kinase B (Akt) is involved in the regulation of neuronal apoptosis. Thirty-six Sprague-Dawley (SD) rats were randomly divided into blank control group, model group and miR-130a low-expression group. miR-130a was determined by quantitative polymerase chain reaction (qPCR), the content of tumor necrosis factor-α (TNF-α), interleukin-6 (IL-6) and IL-10 was detected using the enzyme-linked immunosorbent assay (ELISA) kits, and the neuronal apoptosis level in each group was determined through terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) staining. The neurobehavioral score was significantly lower in model group than that in blank control group (P<0.01), while it was significantly higher in miR-130a low-expression group than that in model group (P<0.01). Compared with blank control group, the model group had obviously increased content of TNF-α and IL-6 (P<0.01), decreased content of IL-10 (P<0.01), more apoptotic neurons (P<0.01), higher expression of caspase-3 (P<0.01), and obviously lower Bcl-2/Bax (P<0.01). Moreover, expression of phosphorylated (p)-PTEN, PI3K and p-Akt in brain tissues was remarkably lower in the model group than those in the blank control group (P<0.01). The expression level of miR-130a in brain tissues of CI rats is significantly increased. miR-130a promotes the release of inflammatory factors and facilitates neuronal apoptosis through suppressing the PTEN/PI3K/Akt signaling pathway.
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Most cited references18
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Intravenous Bone Marrow Stem Cell Grafts Preferentially Migrate to Spleen and Abrogate Chronic Inflammation in Stroke
Sandra Acosta, Naoki Tajiri, Jaclyn Hoover … (2015)
Background and Purpose— Adult stem cell therapy is an experimental stroke treatment. Here, we assessed homing and anti-inflammatory effects of bone marrow stromal cells (hBMSCs) in chronic stroke. Methods— At 60 days post stroke, adult Sprague–Dawley rats received intravenous hBMSCs (4×106 labeled or nonlabeled cells) or vehicle (saline). A sham surgery group served as additional control. In vivo imaging was conducted between 1 hour and 11 days post transplantation, followed by histological examination. Results— Labeled hBMSCs migrated to spleen which emitted significantly higher fluorescent signal across all time points, especially during the first hour, and were modestly detected in the head region at the 12 hours and 11 days, compared with nonlabeled hBMSCs and vehicle-infused stroke animals, or sham (P<0.05). At 11 days post transplantation, ex vivo imaging confirmed preferential hBMSC migration to the spleen over the brain. Hematoxylin and eosin staining revealed significant 15% and 30% reductions in striatal infarct and peri-infarct area, and a trend of rescue against neuronal loss in the hippocampus. Unbiased stereology showed significant 75% and 60% decrements in major histocompatibility complex II–activated inflammatory cells in gray and white matter, and a 43% diminution in tumor necrosis factor-α cell density in the spleen of transplanted stroke animals compared with vehicle-infused stroke animals (P<0.05). Human antigen immunostaining revealed 0.03% hBMSCs survived in spleen and only 0.0007% in brain. MSC migration to spleen, but not brain, inversely correlated with reduced infarct, peri-infarct, and inflammation. Conclusions— hBMSC transplantation is therapeutic in chronic stroke possibly by abrogating the inflammation-plagued secondary cell death.
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Reperfusion therapy in acute ischemic stroke: dawn of a new era?
Sonu Bhaskar, Peter Stanwell, Dennis Cordato … (2018)
Following the success of recent endovascular trials, endovascular therapy has emerged as an exciting addition to the arsenal of clinical management of patients with acute ischemic stroke (AIS). In this paper, we present an extensive overview of intravenous and endovascular reperfusion strategies, recent advances in AIS neurointervention, limitations of various treatment paradigms, and provide insights on imaging-guided reperfusion therapies. A roadmap for imaging guided reperfusion treatment workflow in AIS is also proposed. Both systemic thrombolysis and endovascular treatment have been incorporated into the standard of care in stroke therapy. Further research on advanced imaging-based approaches to select appropriate patients, may widen the time-window for patient selection and would contribute immensely to early thrombolytic strategies, better recanalization rates, and improved clinical outcomes.
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Involvement of MicroRNAs in the Regulation of Muscle Wasting during Catabolic Conditions*
Ricardo Soares, Stefano Cagnin, Francesco Chemello … (2014)
Introduction Skeletal muscle is a major site of metabolic activity and the most abundant tissue in the human body accounting for almost 50% of the total body mass. Being the largest protein reservoir, muscle serves as a source of amino acids to be utilized for energy production by various organs during catabolic periods (1). For instance, amino acids generated from muscle protein breakdown are utilized by the liver to produce glucose and support acute phase protein synthesis (1). A number of catabolic disease states, including sepsis, burn injury, cancer, AIDS, diabetes, heart and renal failure are characterized by muscle wasting, mainly reflecting increased breakdown of myofibrillar proteins. Loss of muscle proteins results in muscle atrophy and weakness that have significant clinical consequences resulting in bad prognosis and life expectancy. Conversely, some forms of physical activity such as endurance exercise prevent a decrease of skeletal muscle mass and induces clinical benefits for patients (2 – 4). However, the molecular mechanisms and the signaling pathways that control muscle strength and size are largely unknown and only recently have some of them been unraveled. Skeletal muscle mass is sustained by a balance between accumulation of newly synthesized proteins and degradation of existing proteins. Different pathways regulate the balance of these two processes and affect muscle performance (5). This view was recently advanced by the finding that muscle atrophy requires a transcriptional regulation of a subset of key genes. The analysis of the gene expression profiles of muscle collected from different atrophic models (i.e. diabetes, cancer cachexia, chronic renal failure, fasting, and denervation) led to the identification of a subset of genes that are commonly up- or down-regulated in muscle atrophy. Because all these diseases share muscle atrophy as a common trait, these genes were called atrophy-related genes or atrogenes and are believed to regulate the loss of muscle components (6). These findings indicate that muscle atrophy is an active process controlled by specific signaling pathways and transcriptional programs. In fact, several transcription factors have been found to regulate muscle atrophy (7 – 12). Considering the complexity of the atrophic program, other levels of regulation may be involved. MicroRNAs (miRNAs) 5 are endogenous ∼22 nucleotide small noncoding RNAs that control gene expression by targeting mRNAs and triggering either translation repression or RNA degradation. These noncoding RNAs bind to the 3′UTR of the target mRNA, and the specificity of the binding is dictated mainly by the seed region of the miRNA (from nucleotide 2 to 8). Hundreds of target genes, often belonging to the same pathway, are regulated by the same miRNA. To increase the complexity of this post-transcriptional regulation several miRNAs can target the same transcript (13). miRNAs are required for several biological processes and complete lack of miRNAs is incompatible with life (14, 15). Interestingly, many miRNAs are expressed in a tissue-specific manner (16). This is the case of several miRNA specifically expressed in cardiac and skeletal muscles and, therefore, named myo-miRs (16 – 18). In striated muscles, miRNAs are involved in physiological processes such as myogenesis, fiber-type switch, and regeneration (19 – 21). Consequently, deregulation of the miRNA expression levels is associated with pathological conditions (22, 23). For instance, inhibition of miRNA-206 in SOD1G93A transgenic mice induced severe atrophy (24). However, the involvement of miRNAs in skeletal muscle atrophy is largely unknown. To determine which miRNAs are relevant for the atrophic process, we performed miRNA expression profiling in muscles from different atrophic models including fasting, denervation, streptozotocin-induced diabetes, and cancer cachexia. We checked whether there was a common signature of miRNA expression in different atrophying conditions and found that each catabolic situation is associated to a peculiar set of deregulated miRNAs. We further focused on atrophy induced by denervation and identified the two most up-regulated miRNAs, miRNA-206 and miRNA-21. In vivo loss- and gain-of-function experiments defined the functional relevance of these two miRNAs for muscle mass regulation. Finally, by combining gene expression profiling and bioinformatic prediction tools we identified and characterized two downstream targets of these atrophy miRNAs. All together, these results start to unravel, in vivo, the role of miRNAs in the atrophic process and open the possibility to manipulate microRNA expression as a novel therapeutic approach to counteract muscle loss in catabolic conditions. EXPERIMENTAL PROCEDURES Animal Models and Surgical Procedures All experimental procedures were approved and authorized by the Italian Ministry of Health. Mice were housed in individual cages in an environmentally controlled room (23 °C, 12-h light-dark cycle) and provided food and water ad libitum. Four different models of skeletal muscle wasting were used: starvation, denervation, diabetes, and cancer cachexia. Starvation, denervation, and diabetes were performed on adult, 2-month-old, CD1 mice, whereas cancer cachexia was induced on 7-week-old BALB/c mice. Fasting was performed as previously described, by removing chow with free access to water. Denervation experiments were performed by cutting the sciatic nerve of one limb, whereas the other was used as control. Mice were sacrificed after 3, 7, or 14 days after denervation. Diabetes was induced by one single acute intraperitoneal (IP) injection of 180 mg/kg of streptozotocin (Sigma S-0130). To induce cancer-associated cachexia, a 0.5-mm3 solid fragment of colon carcinoma (C26, obtained from the National Cancer Institute) was subcutaneously implanted in the back of 7-week-old BALB/c mice (Charles River). Mice were killed 14 days after tumor injection, when the tumor bearing mice had a body weight loss of ∼25% compared with control mice, and hindlimb muscles were removed. For this model mice were housed individually in the Animal Care Facility at the Unit of Histology and Medical Embryology (Rome). After Animal Death Muscles Were Collected and Immediately Frozen in Liquid Nitrogen for Further Analysis Tibialis anterior (TA) muscles of adult CD1 male mice (28–34 g) were transfected as described previously (9, 25). Briefly, plasmid DNA was injected along the muscle length. Electric pulses were then applied by two stainless steel spatula electrodes with the Electro Square Porator (ECM 830, BTX). Transfected muscles were collected 7, 10, or 14 days after electroporation. Cell Culture For in vitro experiments we used C2C12 (mouse myoblast) cells, bought from ATCC. Cells were maintained in culture with DMEM (Dulbecco's modified Eagle's medium) containing 10% FBS and 1% penicillin-streptomycin at 37 °C and 5% CO2. C2C12 cells were maintained in proliferation and were not differentiated into myotubes. Transfection was performed using the Lipofectamine 2000 reagent (Invitrogen 11668-027) according to the manufacturer's protocol. One day prior to the transfection, cells were plated onto 6-well plates at a density of 100,000 cells per well. Cells were transfected with a total of 4 μg of plasmidic DNA per well (6-well plate). Cells were used 24 or 48 h after transfection, according to the experiments. RNA and miRNAs Purification For the microarray experiments mRNA and miRNAs were isolated from frozen gastrocnemius muscles. miRNAs/mRNAs isolation was achieved by an initial purification with TRIzol (Invitrogen) followed by subsequent purification and fractionation with the Purelink miRNA Isolation Kit (Invitrogen number K1570-01). Total and small RNA were quantified using the ND-1000 spectrophotometer (Nanodrop), whereas RNA integrity and content of microRNAs (%) in each sample were assessed by capillary electrophoresis using the RNA 6000 Nano LabChip and the Small RNA Nano LabChip, respectively, using the Agilent Bioanalyzer 2100 (Agilent Technologies). Only total RNA samples with RNA integrity number values >6, and the percentage of miRNA 200 nucleotides) samples were labeled using the Amino Allyl MessageAmpTM II aRNA Amplification Kit (Ambion, Austin, TX), in accordance with the manufacturer's instructions. Briefly, first strand synthesis with an engineered reverse transcriptase should produce virtually full-length cDNA, which is the best way to ensure reproducible microarray results. The use of a modified oligo(dT) primer bearing a T7 promoter (27) allows the next amplification steps: after second strand synthesis and clean-up the cDNA becomes a template for in vitro transcription with T7 RNA polymerase. About 5 μg of aminoallyl-labeled aRNA were coupled with Cy5 or Cy3 dyes (GE Healthcare) and purified on a column (Ambion). Labeled targets (atrophic and control muscle) were mixed and ethanol precipitated. After dissolving the pellet in 120 μl of hybridization buffer (5× SSC, 0.1% SDS, 25% formamide), samples were denatured at 90 °C for 2 min and added to the microarrays. Prehybridization was performed overnight at 48 °C in the presence of 5× SSC, 5× Denhardt, 0.1% SDS, 100 ng/μl of single-stranded DNA. Competitive hybridizations were carried on for 48 h at 46 °C in an ArrayBooster microarray incubator (Advalytix), followed by a series of post-hybridization washings (28). Slides were scanned on a GSI Lumonics LITE dual confocal laser scanner (PerkinElmer Life Sciences) and QuantArray software (GSI Lumonics) was used for image analysis. Raw data are available on GEO database using accession number GSE52676. Statistical Analysis of miRNA and Gene Expression Data Interarray normalization of expression levels performed with loessM+GPA for miRNA experiments and a within-array-normalization, using the lowess method, was performed by using the MIDAW tool for gene expression profiling (29). Normalization function was applied to miRNA expression data of all experiments and then values of spot replicates within arrays were averaged. Cluster analysis and profile similarity searching were performed with tMev that is part of the TM4 Microarray Software Suite. In particular, hierarchical cluster analysis was performed with a Euclidean distance coefficient as distance measure with complete linkage. One-way analysis of variance was used to identify miRNAs differentially expressed (p value ≤ 0.05 based on 1,000 permutation). Pathway analysis was performed using the WEB-based GEne SeT AnaLysis Toolkit (30) and KEGG database on differentially expressed genes of denervated muscles compared with controls through SAM analysis (31). Benjamini and Hochberg multiple test adjustment was used to the determine pathway significance and the top 10 pathways were considered. Identification of miRNA Target Genes One of the most important and difficult steps in miRNA investigations is the identification of their target genes to understand the mechanism by which miRNAs can modulate a certain biological function. We have first recovered potential mRNA targets calculated by the PITA algorithm (32). Hundreds of target genes for any single miRNA have been predicted by this analysis. To limit the fraction of false positives we have analyzed the functional anticorrelation between miRNAs and mRNA expression levels (e.g. miRNA is up-regulated and corresponding mRNA target is down-regulated) (Fig. 5 A) by searching specific profiles of miRNAs and mRNAs through the template matching algorithm (33). Gene ontology classification of miRNA-206 and -21 targets was performed using the DAVID web tool (34). Validation of the Microarray Results miRNA expression profiles were validated using the TaqMan® MicroRNA Assays (Applied Biosystems). Retrotranscription of each specific miRNA was accomplished using the TaqMan MicroRNA Reverse Transcription Kit (Applied Biosystems) according to the manufacturer's instructions. RT-PCR was then preformed using the TaqMan Universal PCR Master Mix and the specific primers from TaqMan MicroRNA Assay specific for each miRNA. miRNAs expression was normalized with the U6 small nuclear RNA (U6 snRNA) (Applied Biosystems, assay ID 001973). Studied miRNA were miRNA-21 (Applied Biosystems, assay ID 000397), miRNA-206 (Applied Biosystems, assay ID 000510), and miRNA-133b (Applied Biosystems, assay ID 002247). Gene expression analysis was performed by qRT-PCR. Complementary DNA was generated using Superscript III Reverse transcriptase (Invitrogen number 18080044). cDNA was amplified, using Power SYBR Green PCR Master Mix (Applied Biosystems number 4367659), with an ABI Prism 7900HT (Applied Biosystem) thermocycler. Gapdh and Actin were used as a reference to normalize expression data. The sequence of the oligonucleotide primers used is shown in Table 1. TABLE 1 List of primers used for real time experiments Primer Sequence GAPDH-forward 5′-CACCATCTTCCAGGAGCGAG-3′ GAPDH-reverse 5′-CCTTCTCCATGGTGGTGAAGAC-3′ Pan-Actin-forward 5′-CTGGCTCCTAGCACCATGAAGAT-3′ Pan-Actin-reverse 5′-GGTGGACAGTGAGGCCAGGAT-3′ Yy1-forward 5′-TGAGAAAGCATCTGCACACC-3′ Yy1-reverse 5′-CGCAAATTGAAGTCCAGTGA-3′ Eif4e3-forward 5′-AACATCCCTCCTGTGACCAG-3′ Eif4e3-reverse 5′-TCCAATGGTCGCTAACAACA-3′ Pdcd10-forward 5′-GGGCACTTGAACACCAAAAG-3′ Pdcd10-reverse 5′-CAGGCCACAGTTTTGAAGGT-3′ Cloning and Plasmids Transfection and electroporation experiments were preformed with the following constructs. pmiRZIP lentivector (ZIP NULL), pmiRZIP lentivector anti-miRNA-206 (ZIP-206), and pmiRZIP lentivector anti-miRNA-21(ZIP-21) were acquired from System Bioscience. pMIR206-Luc and pMIR21-Luc were acquired from Signosis BioSignal. Renilla Null and Renilla TK were acquired from Promega (pRL-null number E2271, pRL-TK number E2241). The mature sequences of miRNA-206 and miRNA-21 were cloned into the BLOCK-iT Pol II miR RNAi Expression Vector Kit with EmGFP (Invitrogen number K4936-00). The oligos used are shown in Table 2. As negative control the pcDNA6.2-GW/EmGFP-miR-neg control was used, according to manufacturer's instructions. TABLE 2 List of primers used in cloning experiments Name Sequence miRNA-206, top 5′-TGCTGTGGAATGTAAGGAAGTGTGTGGGTTTTGGCCACTGACTGACCCACACACCC TACATTCA-3 miRNA-206, bottom 5′-CCTGTGAATGTAGGGTGTGTGGGTCAGTCAGTGGCCAAAACCCACACACTTCCTTACATTCCAC-3 miRNA-21, top 5′-TGCTGTAGCTTATCAGACTGATGTTGAGTTTTGGCCACTGACTGACTCAACATCTCTGATAAGCTA-3′ miRNA-21, bottom 5′-CCTGTAGCTTATCAGAGATGTTGAGTCAGTCAGTGGCCAAAACTCAACATCAGTCTGATAAGCTAC-3′ Yy1-3′UTR-forward 5′-GCCTCTTCAGGAGTGTGATTG-3′ Yy1-3′UTR-reverse 5′-CAATTTCTGGGAGGCTCAAG-3′ Eif4e3-3′UTR-forward 5′-TCTGCCATCGTATCACTTGC-3′ Eif4e3-3′UTR-reverse 5′-GCCTCTTACGCTCTGACCAC-3′ Pdcd10-3′UTR-forward 5′- ACTAGTCCAGGATGTTGAATGGGATT-3′ Pdcd10-3′UTR-reverse 5′- GCGGCCGCAAGTAAAGAAATGTTTAACA-3′ PolK-3′UTR-forward 5′- ACTAGTCCTTTAAGGAAGACAAGTGCAA-3′ PolK-3′UTR-reverse 5′- GCGGCCGCCAACAAAAATAAACTTCAGATGGAA-3′ The 3′UTR of the different analyzed genes was cloned from muscle cDNA into the pMIR-LUC vector (Signosis BioSignal). The sequence of the oligonucleotide primers used to clone the 3′UTR of each gene are shown in Table 2. All constructs were sequenced to confirm the insert and absence of mutation. To mutate the 3′UTR of YY1 and eIF4E3 the QuikChangeII site-directed Mutagenesis Kit (Stratagene number 200524) was used according to the manufacturer's instructions. The primers used for the mutagenesis are shown in Table 3. All mutated constructs were sequenced to confirm the presence of the desired mutations and the absence of other unspecific mutations. TABLE 3 List of primers used for the mutagenesis experiments Primer Sequence Eif4e3_mut-206_forw 5′-AAAGTCAGGGGCCTCCACTTGAAGCGCTAAACAGGAAGCCAAATTA-3′ Eif4e3_mut-206_rev 5′-TAATTTGGCTTCCTGTTTAGCGCTTCAAGTGGAGGCCCCTGACTTT-3′ Eif4e3_mut-21_forw 5′-GCCTAGCAAAACCCTTTTTCTGCTGCATTGTTGTGACACTTCCCTGCA-3′ Eif4e3_mut-21_rev 5′-TGCAGGGAAGTGTCACAACAATGCAGCAGAAAAAGGGTTTTGCTAGGC-3′ Yy1_mut-1_forw 5′-GTGCATATTGTACACTTTTTGGGGATCTTATTAGTAATGCTGTGTGATTTTCTGGA-3′ Yy1_mut-1_rev 5′-TCCAGAAAATCACACAGCATTACTAATAAGATCCCCAAAAAGTGTACAATATGCAC-3′ Yy1_mut-2_forw 5′-GCTGTGTGATTTTCTGGAGGTTGATCGCTGTGCTTGCGGTAGATTTTCTTT-3′ Yy1_mut-2_rev 5′-AAAGAAAATCTACCGCAAGCACAGCGATCAACCTCCAGAAAATCACACAGC-3′ Cross-sectional Area Measurements Cross-sectional area of GFP-positive transfected fibers was measured as described previously (8) and compared with the surrounding nontransfected myofibers (control). The pictures of the transfected muscles were taken at ×20 magnification with a fluorescence microscope (Zeiss AxioImager.Z1). Fiber cross-sectional areas were measured using IMAGE software (Scion). All data are expressed as the mean ± S.E. Comparison were made by using the Student's t test, with p 3 muscles per condition. *, p 3 muscles per condition. *, p 3 muscles per condition. *, p 3 muscles per condition. *, p 3 muscles per condition. **, p 3 muscles per condition. J, miRNA-206 and -21 luciferase sensors were inhibited in denervated muscles. Data are shown as mean ± S.E. n > 4 muscles per condition; *, p 1.2 are represented for each atrophic condition (blue is for down-regulation and yellow is for up-regulation) Quantitative RT-PCR confirmed the up-regulation of miRNA-206 and miRNA-21 after denervation even if their pattern of expression was slightly different (Fig. 1, D and E). MicroRNA-206 was also induced by fasting but not by diabetes (Fig. 1, F and H), whereas miRNA-21 was suppressed by fasting and unchanged in diabetes (Fig. 1, G and I). Due to the important induction of both of these microRNAs in denervated muscles we focused our attention on this model of muscle atrophy. To further validate miRNA-206 and -21 up-regulation after denervation we used a luciferase-based assay to monitor the activity of the endogenous miRNAs. We used a vector that contains a binding site specific for each miRNA in the 3′UTR of the luciferase gene. When the miRNA of interest is induced, it binds the luciferase transcript blocking its translation. The miRNA-Luc sensors were transfected into TA muscles of adult mice and then animals were denervated. Luciferase activity of miRNA-206 and miRNA-21 sensors was significantly reduced after 7 days of denervation (Fig. 1 J) confirming the increased activity of these miRNAs after denervation. Because skeletal muscles contain different fiber types that show different metabolic properties, we monitored miRNA expression in denervated fast and slow muscles. miRNA-206 and miRNA-21 were induced in both TA, a glycolytic mitochondrial poor muscle, and soleus, an oxidative mitochondrial rich muscle (Fig. 2, A and B) suggesting that miRNA response is a common feature of atrophying muscles. FIGURE 2. miRNA-206 and -21 are selectively induced in atrophying muscles. A, miRNA-206, and B, miRNA-21 expression levels are increased after 15 days of denervation in both fast and slow muscles. Quantitative RT-PCR analysis on miRNA expression levels in TA and soleus muscles of control and denervated animals. Data are shown as mean ± S.E. n > 3 independent experiments; *, p 3 muscles per condition, *, p 3 muscles per condition; *, p 800 per each muscle, at least 3 muscles per condition were examined; **, p 700 for each muscle, at least 3 muscles per condition were analyzed; **, p 3 independent experiments; *, p 3 muscles per condition; **, p < 0.01. B, eIF4E3 is a target of both miRNA-206 and miRNA-21. Adult TA muscles were transfected and processed as described in A; **, p < 0.01. C, Pdcd10 is a target of both miRNA-206 and miRNA-21. Adult TA muscles were transfected and processed as described in A; **, p < 0.01. D, mutation of the miRNA-206 binding site on the 3′UTR of eIF4E3 partially prevents miRNA-dependent inhibition. C2C12 myoblasts were co-transfected with control or mutated eIF4E3–3′UTR together with scramble or miRNA-206 expressing vector. Luciferase/Renilla ratio was determined 24 h after transfection. Data are shown as mean ± S.E., n = 3 independent experiments; **, p < 0.01; #, p < 0.05. E, mutation of the miRNA-21 binding sites on the 3′UTR of YY1 and eIF4E3 partially prevent miRNA-dependent inhibition. C2C12 myoblasts were co-transfected with the control or mutated version of the 3′UTR of YY1 or eIF4E3 together with scramble or miRNA-21 expressing vector. Luciferase/Renilla ratio was determined 24 h after transfection. Data are shown as mean ± S.E., n = 3 independent experiments; *, p < 0.05; **, p < 0.01; #, p < 0.05; ##, p < 0.01. FIGURE 7. A, over-expression of miRNA-206 or miRNA-21 directly regulates the mRNA of YY1 and eIF4E3. C2C12 myoblasts were transfected with miRNA-206 or miRNA-21 expressing vector. RNA was extracted 48 h after transfection and the expression levels of YY1 and eIF4E3 were analyzed by qRT-PCR. Data are shown as mean ± S.E., n = 3 independent experiments; *, p < 0.05; **, p < 0.01. B, densitometric quantification of the Western blot for YY1 in C2C12 myoblasts over-expressing negative control or miRNA-21. Data are shown as mean ± S.E., n = 3 independent experiments. DISCUSSION Our data demonstrate that miRNA expression is affected by catabolic conditions and modulates the atrophy program after denervation. Moreover, miRNA induction in denervated muscles is delayed when compared with the transcriptional regulation. In fact 3 days after denervation is a time in which muscles are not yet atrophic and are characterized by the peak of mRNAs induction while miRNAs are mainly suppressed. Instead at 7 days of denervation muscles are already atrophic and miRNAs are induced. Altogether these findings suggest that microRNAs are involved in the fine-tuning of an already initiated atrophy program. This concept is also sustained by the minor changes that miRNA-206 and miRNA-21 over-expression and inhibition elicited on myofiber size. A similar observation was achieved in human muscle stem cell (42). Accordingly, miRNAs were globally down-regulated in quiescent compared with proliferating satellite cells. The recent findings that DICER1 and EIF2C/AGO, two essential components of the miRNA machinery, are substrates of the autophagy-lysosome system define a link between autophagy activation and suppression of microRNA maturation (43). Because autophagy is activated at early stages of muscle atrophy (44 – 47) it is possible that suppression of miRNA biogenesis and processing depends on autophagy-dependent degradation of DICER and AGO. However, when we looked at AGO2 expression we could not find a significant down-regulation at 7 days of denervation. Two other studies conducted in different models of cardiac hypertrophy (48) and in several inherited muscular disorders (22) have analyzed the miRNA signature and identified a small subset of miRNAs commonly induced during these diseases. However, it is worth noting that most of the deregulated miRNAs in cardiac hypertrophy or in inherited muscle diseases were specific of each pathological condition. This is in line with our data that did not identify a common signature of miRNA in different models of muscle atrophy. The peculiarity of the miRNA signature for each atrophic condition further confirms the concept of a specific fine-tuning/regulation of an activated atrophy process. This is the first study that compared miRNAs expression in early stages of muscle atrophy and dissected their function, in vivo. Other studies described the changes that occur in diabetes (49), long term denervation/reinnervation (50, 51), and dexamethasone-induced atrophy(52). The decision to focus on denervation was driven by the fact that this model displayed the strongest induction of microRNAs. Moreover, despite the fact that transcriptomes of denervated muscles are well characterized (6, 35, 53), miRNA expression is still unexplored. A few published studies have defined the microRNA profile in long term denervated muscles (1 and 4 months)(50, 51). However, these muscles are in fully established atrophy and therefore, the changes of miRNA profile are mainly related to metabolic adaptations/compensations that follow such long inactivity. Instead, our analyses at early time points unraveled the role of miRNA before and/or during the early stages of atrophy program. This approach allows us to identify miRNA-206 and miRNA-21 as important modulators of muscle loss. miRNA-206 is a muscle-specific microRNA that has been found to be involved in myoblast differentiation (36, 54), muscle regeneration (21), and nerve growth after injury (24). Previous work identified miRNA206 as a critical factor of muscle reinnervation in SOD1G93A transgenic mice, an animal model of amiotrophic lateral sclerosis that is characterized by motor-neuron degeneration. Despite the evident effect of miRNA-206 deletion in SOD1G93A transgenic animals, miRNA-206 knock-out mice did not show any evident phenotype in basal conditions (24). However, a compensation operated by miRNA-1, another muscle-specific miRNA, may account for the lack of phenotype in these knock-out mice. In fact miRNA-206 and miRNA-1 share the same seed region and differ only in 3 nucleotides at the 3′-end region (55), suggesting that they possibly regulate the same set of mRNAs targets and therefore, possibly identical biological processes. By using an electroporation technique, we acutely perturbed the miRNA content of adult muscle minimizing compensatory mechanisms. Our findings are consistent with a recent study that identified miRNA-206 as an important player in myotube size. However, such changes were not confirmed when the authors moved to in vivo experiments (56). The discrepancy of this result with our data could be explained with the level of expression of miRNA-206. In this study the authors used AAV-mediated delivery of miRNA-206 that resulted in a 20- and 100-fold increase of miRNA206 in muscles. Our transfection approach gave a 2–3-fold up-regulation of mature miRNAs mimicking the endogenous induction that occurs in denervated muscles. Therefore, our experimental condition did not saturate the miRNA machinery reducing the possibility of off-target effects. MicroRNA-21 is mainly known for its anti-apoptotic and oncogenic potential (57). miRNA-21 is also important in the cardiovascular system because it is highly up-regulated during cardiac stress, but its actual role is still controversial. Interestingly miRNA-21 was induced in 8 of 10 inherited muscular diseases (22) suggesting that it can be part of the complex mechanisms of muscle wasting. However, when we overexpressed miRNA-21 in innervated muscle we did not induce muscle atrophy suggesting that miRNA-21 requires the co-expression of other microRNAs to fully elicit an atrophic action. The kinetic of miRNA-21 and -206 induction in denervated muscles suggests that miRNA-206 is one of the miRNA-21 partners and that the simultaneous induction of both microRNAs is critical for an optimal down-regulation of important targets controlling muscle mass. In fact miRNA-21 is already up-regulated at 3 days of denervation, a time in which muscles are not atrophic and miRNA-206 is not significantly induced. Importantly, at 7 days both miRNAs are induced and muscles are 25% smaller than controls. This was confirmed by experiments in which miRNA-206 and miRNA-21 were co-expressed leading to an exacerbation of muscle atrophy induced by just miRNA-206 expression. 6 To understand which mechanisms are controlled by these miRNAs we performed an mRNAs expression profile in parallel to the miRNAs expression profile. Among the statistically significant activated pathways in denervated muscles we identified the proteasome, phagosome, protein digestion, and absorption pathways (Table 7), whereas metabolic pathways like the insulin signaling pathway was turned off (Table 8) (58, 59). The microarray analyses identified 4,346 transcripts down-regulated between 3 days of denervation and 7 or 14 days (Fig. 5 B), an opposite profile of that of miRNA-21 and -206. The comparison of down-regulated transcripts with the predicted miRNA targets led to a list of candidates that were further reduced when we selected transcripts regulated by both miRNAs. This strategy led to the identification of YY1, eIF4E3, and PDCD10. Although the function of these genes in atrophy is unclear or unknown, we confirmed their miRNA-dependent suppression in denervation. The YY1 transcription factor is involved in mitochondrial biogenesis in adult muscles (60) through the interaction with PGC1a. Inactivation of YY1 in muscle causes abnormalities of mitochondrial morphology and oxidative function associated with exercise intolerance (61, 62). This finding opens the possibility that miRNA-21, through the regulation of YY1, participates in the regulation of mitochondria biogenesis/functioning. In fact, it is well established that loss of innervation causes a suppression of mitochondrial function and β-oxidative metabolism (6, 63, 64). Therefore, microRNA-dependent suppression of YY1 can contribute to these metabolic adaptations after denervation. TABLE 7 Pathways activated during denervation Activated Proteosome Phagosome MAPK signaling pathway Ribosome p value 1.52E-13 1.03E-09 6.47E-06 8.75E-06 Genes Psmb4 Atp6v0e Flnc Rps18 Psmc1 Itgb1 Cacng1 Rps5 Psmb1 Thbs3 Traf2 Rpl22 Psma7 Atp6v1g1 Relb Rpl23 Psmb5 H2-Eb1 Mknk2 Rps10 Psmd4 Atp6v1e1 Gadd45a Rps9 Psmd2 Tubb6 Rras2 Rpl10a Psmd14 Calr Hspb1 Psma4 Tap1 Rras Psmc4 Rab5a Mapt Tubb2a Tuba1b TABLE 8 Pathways inhibited during denervation Inhibited Metabolic pathways PPAR signaling pathway Amyotrophic lateral sclerosis (ALS) Insulin signaling pathway p value 5.34E-08 7.53E-08 2.20E-06 4.65E-06 Genes Abo Got1 Acadm Casp12 Mapk1 Acadm Hmgcr Cd36 Grin1 Phka1 Adk Ldhb Cpt1b Grin2b Ppargc1a Aldh6a1 Mat2b Cyp27a1 Nos1 Ppp1cb Amd1 Mccc1 Fabp3 Ppp3r1 Ppp1r3c Bpgm Mpst Lpl Slc1a2 Slc2a4 Cs Ndst3 Ppara Socs1 Csl Nnt Rxra Srebf1 Cyp27a1 Nos1 Cyp2c65 Pla2g7 Dbt Prodh Dhcr24 Prodh2 Ephx2 St3gal6 Galt The initiator factor eIF4e3 is one of the most critical factors that regulate ribosome assembly and protein synthesis. Importantly, eukaryotic translation initiation factor 4E-binding protein 1 (eIF4e3-BP1 or simply 4EBP1) is a repressor of protein synthesis and belongs to the group of atrophy-related genes(59). The transcriptional-dependent up-regulation of 4EBP1 and the miRNA-dependent inhibition of eIF4E3 suggest a coordinated action to finely tune protein synthesis in denervated muscles. Moreover, our finding that two miRNAs control eIF4E3 expression is consistent with the concept that miRNA-206 and miRNA-21 synergistically act to slow down the rate of protein synthesis. We cannot exclude that other targets are involved in the regulation of muscle mass. Especially the data that miRNA-206 alone is sufficient to induce atrophy may support this concept. Indeed, the potential miRNA-206 targets include factors, such as Smad1, Runx1, JunD, and Rheb, that have been described to affect muscle mass. The contribution of miRNA-206 in the regulation of these targets will be evaluated in the future. In conclusion, our data propose the novel hypothesis that miRNAs act as important players in the modulation of the atrophy program that controls critical components of mitochondria function and protein synthesis.
Author and article information
Journal
Journal ID (nlm-ta): Exp Ther Med
Journal ID (iso-abbrev): Exp Ther Med
Journal ID (publisher-id): ETM
Title:
Experimental and Therapeutic Medicine
Publisher:
D.A. Spandidos
ISSN
(Print):
1792-0981
ISSN
(Electronic):
1792-1015
Publication date
(Print):
March
2020
Publication date
(Electronic):
03
January
2020
Publication date PMC-release: 03
January
2020
Volume: 19
Issue: 3
Pages: 2119-2126
Affiliations
Author notes
Correspondence to: Dr Shuhua Gui, Department of Neurology, The Third People's Hospital of Wuxi, 585
Xinyuanbei Road, Liangxi, Wuxi, Jiangsu 214041, P.R. China, E-mail:
ikdps827@
123456163.com
Article
Publisher ID: ETM-0-0-8415DOI: 10.3892/etm.2020.8415
PMC ID: 7027342
PubMed ID: 32104274
SO-VID: 1d09109d-7c2a-4265-8539-34be5193cb84
Copyright © Copyright: © Wang et al.
License:
This is an open access article distributed under the terms of the Creative Commons Attribution-NonCommercial-NoDerivs License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non-commercial and no modifications or adaptations are made.
History
Date
received
: 08
October
2019
Date
accepted
: 04
December
2019
Categories
Subject:
Articles
ScienceOpen disciplines: Medicine
Data availability:
ScienceOpen disciplines: Medicine