- Open Access
Splicing-dependent expression of microRNAs of mirtron origin in human digestive and excretory system cancer cells
© Butkytė et al. 2016
- Received: 9 February 2016
- Accepted: 18 March 2016
- Published: 25 March 2016
An abundant class of intronic microRNAs (miRNAs) undergoes atypical Drosha-independent biogenesis in which the spliceosome governs the excision of hairpin miRNA precursors, called mirtrons. Although nearly 500 splicing-dependent miRNA candidates have been recently predicted via bioinformatic analysis of human RNA-Seq datasets, only a few of them have been experimentally validated. The detailed mechanism of miRNA processing by the splicing machinery and the roles of mirtronic miRNAs in cancer are yet to be uncovered.
We experimentally examined whether biogenesis of certain miRNAs is under a splicing control by analyzing their expression levels in response to alterations in the 5′- and 3′-splice sites of a series of intron-containing minigenes carrying appropriate miRNAs. The expression levels of the miRNAs processed from mirtrons were determined by quantitative real-time PCR in five digestive tract (pancreas PANC-1, SU.86.86, T3M4, stomach KATOIII, colon HCT116) and two excretory system (kidney CaKi-1, 786-O) carcinoma cell lines as well as in pancreatic, stomach, and colorectal tumors. Transiently expressed SRSF1 and SRSF2 splicing factors were quantified by western blotting in the nuclear fractions of HCT116 cells.
We found that biogenesis of the human hsa-miR-1227-3p, hsa-miR-1229-3p, and hsa-miR-1236-3p is splicing-dependent; therefore, these miRNAs can be assigned to the class of miRNAs processed by a non-canonical mirtron pathway. The expression analysis revealed a differential regulation of human mirtronic miRNAs in various cancer cell lines and tumors. In particular, hsa-miR-1229-3p is selectively upregulated in the pancreatic and stomach cancer cell lines derived from metastatic sites. Compared with the healthy controls, the expression of hsa-miR-1226-3p was significantly higher in stomach tumors but extensively downregulated in colorectal tumors. Furthermore, we provided evidence that overexpression of SRSF1 or SRSF2 can upregulate the processing of individual mirtronic miRNAs in HCT116 cells.
An interplay of different splicing factors, such as SRSF1 or SRSF2, may alter the levels of miRNAs of mirtron origin in a cell. Our findings underline the specific expression profiles of mirtronic miRNAs in colorectal, stomach, and pancreatic cancer.
- Post-transcriptional RNA processing
- Splicing factor
- Human cancer
All multi-exon human genes undergo constitutive and alternative splicing, a very precise process which is crucial for regulation of gene expression and generation of proteomic and functional diversity . In the cell, RNA splicing takes place in the nucleus within a large macromolecular complex, the spliceosome, which consists of five small nuclear ribonucleoproteins (snRNPs) and over a few hundred auxiliary proteins . Somatic mutations or dramatic alterations in the amount of splicing factors under pathologic conditions, particularly in human cancers, have been observed, thus resulting in abnormal expression of tumor suppressors or oncogenes [3–5]. Furthermore, various types of cancer have been linked to dysregulation of microRNAs (miRNAs), short non-coding RNAs that post-transcriptionally regulate gene expression in mammals . Although the majority of miRNA genes possess their own internal promoters that are regulated by different transcription and epigenetic factors, approximately one third of them are processed from introns of protein-coding or long non-coding RNAs genes . That provides means for coupled regulation of miRNA maturation and mRNA splicing. However, the spatiotemporal control of miRNA expression by splicing factors has not been elucidated.
Canonical pri-miRNA is cleaved by Drosha/DGCR8 complex to form pre-miRNA hairpin, which is recognized by Exportin-5 and transferred to the cytoplasm. After processing by Dicer ribonuclease, the guide miRNA strand of mature miRNA/miRNA* duplex is incorporated into a RNA-induced silencing complex (RISC). An alternative pathway of miRNA biogenesis was lately described in eukaryotes [8, 9]. Short introns containing miRNAs, termed mirtrons, are spliced and debranched into pre-miRNA hairpin mimics that appear to bypass Drosha cleavage (Additional file 1: Figure S1). Debranched mirtrons enter the canonical miRNA pathway during nuclear export. Mirtrons were originally described in flies and worms [10, 11], but similar loci (i.e., short hairpin introns associated with small RNA reads extending to intronic termini) were later predicted in a range of plants and mammalians, including humans [12–14]. Based on the nature of terminal overhangs, four distinct mirtron subtypes are recognized: conventional (hairpin without tails), 3′-tailed, 5′-tailed, and two-tailed .
Despite the fact that the first splicing-dependent human mirtrons were annotated in 2007  and a large number of splicing-derived miRNAs have been predicted by bioinformatic analysis of deep sequencing data [12, 14], only two of them, hsa-miR-877-5p and hsa-miR-1226-3p, were experimentally proven to be directly affected by splicing [16, 17]. 478 mirtron candidates predicted by Wen et al.  could cover a significant part of the total human miRNA population (2588 mature miRNA entries according to miRBase release 21 ), but little is still known about their impact on the expression of disease-related genes. It is currently shown that the mirtronic miRNA hsa-miR-1226-3p may function as an oncosuppressor by downregulating the mucin 1 oncoprotein and the hsa-miR-1226-3p level is decreased in breast cancer cells . hsa-miR-1236-3p processed from a putative mirtron downregulates alpha fetoprotein, thus leading to the inhibition of the PI3K/Akt pathway in hepatoma cancer cell lines , and negatively regulates the vascular endothelial growth factor receptor VEGFR-3 during inflammatory lymphangiogenesis . In addition, a miR-1236-3p antisense oligonucleotide inhibits the glioma tumor cell growth and proliferation . Overexpression of hsa-miR-877-5p was observed in metastatic melanoma  and endometrial serous adenocarcinomas , but no biological function was determined. Yet, very little is known about the expression profiles of mirtron-derived miRNAs in tumors or cancer cells. There is a high possibility that the expression levels of splicing factors can not only affect alternative pre-mRNA splicing but cause changes in mirtronic miRNA expression as well.
In this study, we examined eight putative mirtrons and provided experimental evidence that human hsa-miR-1227-3p, hsa-miR-1229-3p, and hsa-miR-1236-3p could be assigned to the class of mirtronic miRNAs. Digestive and excretory system cancer cell lines, as well as digestive system tumors tissues, display varying expression profiles of the previously identified hsa-miR-1226-3p and the two newly validated mirtronic miRNAs. Finally, we found that overexpression of well-known splicing factors SRSF1 and SRSF2 increased the abundance of some mirtron-derived miRNAs in colorectal HCT116 cancer cells.
Experimental validation of new splicing-dependent human mirtronic miRNAs
Alterations of miRNA expression level in cells transfected with native (WT) or splicing-deficient (MUT) minigenes
WT minigene linea
MUT minigene linea
Hosted on conventional mirtron
81 ± 36
1.5 ± 0.3
405 ± 204
24 ± 8
1633 ± 106
4.1 ± 2.1
747 ± 217
2.6 ± 0.5
1772 ± 1289
425 ± 322
Hosted on 5′-tailed mirtron
27 ± 13
47 ± 29
875 ± 162
1541 ± 576
Hosted on 3′-tailed mirtron
9.1 ± 1.7
13 ± 0.3
20 ± 12
57 ± 38
Differential expression of miRNAs processed from conventional mirtrons in cancer cell lines
Differential expression of mirtronic miRNAs in pancreas, kidney, colon, and stomach cancer cell lines
miRNAs of mirtron origin
−2.8 ± 1.8 a
−1.7 ± 0.3 a
−2.8 ± 0.9 a
−1.4 ± 0.2
−1.2 ± 0.6
−3.4 ± 0.9 a
−2.1 ± 0.4 a
+5.0 ± 2.7 b
+3.8 ± 0.6 b
+2.8 ± 0.9 b
−2.2 ± 0.4 a
−2.6 ± 0.4 a
+2.1 ± 0.3 b
+1.9 ± 0.4 b
+1.3 ± 0.2
−4.9 ± 2.1 a
−3.9 ± 1.6 a
−1.2 ± 0.6
+1.3 ± 0.7
1.0 ± 0.8
−4.6 ± 2.2 a
−2.6 ± 0.7 a
−1.1 ± 0.2
−1.8 ± 0.2 a
−1.1 ± 0.2
−2.7 ± 0.8 a
−2.2 ± 0.8 a
−1.3 ± 0.2
1.0 ± 0.7
−4.2 ± 0.3 a
−1.9 ± 0.3 a
−2.0 ± 0.4 a
+2.7 ± 0.3 b
−2.2 ± 0.3 a
+1.3 ± 0.0
The expression pattern of the tested mirtron-derived miRNAs revealed a unique miRNA signature of individual cancer cell lines (Fig. 2). The abundance of two of them, hsa-miR-1226-3p and hsa-miR-1227-3p, was reduced two- to five-fold in all cancer cell lines relative to the embryonic kidney HEK 293A cells. The highest downregulation was observed in the kidney carcinoma CaKi-1 and 786-O cells. In contrast, hsa-miR-1229-3p and hsa-miR-1236-3p showed significant variations among cancer cell lines and have a potential to be exploited for profiling specific cancer cell types. Correlations were found between the hsa-miR-1229-3p expression pattern and tissue origin — a steep increase in the mirtronic miRNA abundance was detected in three lines (SU.86.86, T3M4, and KATOIII) derived from metastatic sites as opposed to those derived from primary tumors. In particular, the PANC-1 cells (established from pancreatic carcinoma of ductal origin) showed a 14-fold reduction of this mirtronic miRNA as compared to the SU.86.86 cells (derived from pancreatic ductal carcinoma metastatic site in the liver). Moreover, hierarchical clustering based on relative expression variations for mirtronic miRNAs identified the majority of the cell lines derived from metastatic sites as a unique group (Fig. 2). However, further studies are required to ascertain whether hsa-miR-1229-3p is selectively upregulated in metastatic tumor tissues.
The levels of canonical hsa-miR-1238-3p expression in the majority of cancer cells, with the exception of SU.86.86 and HCT116, were similar to embryonic HEK 293A cells.
Expression profiling of mirtronic miRNAs in pancreatic, colorectal, and stomach tumors
To further explore the impact of mirtronic miRNAs on human cancer development, we analyzed whether mirtron-derived miRNAs are differentially expressed in digestive system tumors (pancreatic, colorectal, and stomach) as compared to normal solid tissue samples of the same organ.
Expression of mirtronic miRNAs in colorectal, stomach, and pancreatic cancerous tissues compared to healthy tissues
miRNAs of mirtron origin
Effect of splicing factors SRSF1 and SRSF2 on biogenesis of mirtronic miRNAs
Despite a considerable progress in the study of biogenesis of mirtron-derived miRNAs, the factors responsible for mirtronic intron splicing are not known. Commonly, the efficiency of the macromolecular spliceosome complex is controlled by associated protein cofactors called splicing factors (SFs). However, it is not known whether the splicing proteins directly affect the biogenesis of mirtronic miRNAs in human cells. To evaluate the regulatory function of specific SF in maturation of mirtron-derived miRNAs, we overexpressed two essential sequence-specific SFs of serine-arginine-rich (SR) protein family, SRSF1 (also known as SF2/ASF) and the SRSF2 (also known as SC35), in human HCT116 cells and measured cellular levels of three splicing-dependent miRNAs, hsa-miR-1226-3p, hsa-miR-1227-3p, and hsa-miR-1229-3p. These prototypical SR proteins play an important role in the regulation of both constitutive and alternative pre-mRNA splicing . In addition, SRSF1 is also involved in pri-miRNA processing .
Change of mirtronic miRNA expression levels in response to the overexpression of splicing factors
Overexpressed splicing factor
Change of mirtronic miRNA expression levels, folds (p value)
SRSF1 (3.2 ± 0.6-fold)
1.5 ± 0.6 (p = 0.23)
−1.1 ± 0.4 (p = 0.47)
1.7 ± 0.4a (p = 0.05)
SRSF2 (1.6 ± 0.5-fold)
1.5 ± 0.8 (p = 0.15)
1.9 ± 0.7a (p = 0.008)
1.4 ± 0.3a (p = 0.008)
Overall, our results lead to a conclusion that altered expression of a particular SF may affect the levels of mirtronic miRNAs in the cell.
To date, nearly 500 human miRNA-containing mirtrons have been predicted by bioinformatic analysis of deep sequencing data . Nevertheless, their dependence on splicing was experimentally validated only for two human miRNAs located in conventional mirtrons, hsa-miR-877-5p and hsa-miR-1226-3p [16, 17]. Moreover, there is no convincing evidence for the participation of splicing machinery in biogenesis of miRNAs processed from 5′-tailed mirtrons, the most abundant subtype accounting for 86 % of all predicted mirtrons . Likewise, for 3′-tailed mirtronic miRNAs, the splicing dependency was determined only in Drosophila . Therefore, direct experimental evidence is necessary to confirm assignment of miRNAs to the mirtron pathway. In the present study, we experimentally proved that hsa-miR-1227-3p, hsa-miR-1229-3p, and hsa-miR-1236-3p derived from conventional mirtrons are processed by the splicing machinery. Meanwhile, splicing was not strictly required for the maturation of hsa-miR-1238-3p, previously annotated as mirtron-encoded miRNA [12, 15]. Indeed, Wen et al. recently reclassified the hsa-mir-1238 mirtron in miRBase based on more stringent bioinformatic criteria applied for mirtron evaluation . Despite successful experimental validation of mirtronic miRNAs derived from conventional mirtrons, analysis of the annotated 5′-tailed mirtrons containing hsa-miR-3064-5p and hsa-miR-6515-5p or 3′-tailed mirtrons with hsa-miR-3940-5p and hsa-miR-6850-5p revealed no splicing requirement for the biogenesis of the intron-derived miRNAs. Thus, it is still an open question whether 5′- or 3′-tailed mirtrons actually exist in mammals. Overall, five out of nine tested human mirtrons generate miRNA irrespective of splicing. Similarly, several groups reported that only two of three  and two of four  predicted mirtronic miRNA candidates mature via a mirtron biogenesis pathway. These observations highlight the relevance of experimental validation of predicted mirtrons.
Mirtron-encoded miRNAs were predominantly found to occur in young, recently emerged genes . Despite their relatively short evolution, the newly evolved miRNAs are incorporated into beneficial regulatory networks . Babiarz et al. suggested that mirtronic miRNAs play important functional roles in post-mitotic neurons in the mammalian brain ; however, very little is known about their expression in cancerous cells. In this study, cell line-specific profiles of miRNAs processed from conventional mirtrons, hsa-miR-1226-3p, hsa-miR-1227-3p, hsa-miR-1229-3p, and hsa-miR-1236-3p, in seven various pancreatic, kidney, colorectal, and stomach cancer cell lines were identified (Table 2). The miRNA variability in cancer cell lines was determined by changes in abundance of two mirtronic miRNAs, hsa-miR-1229-3p and hsa-miR-1236-3p, while the expression levels of hsa-miR-1226-3p and hsa-miR-1227-3p declined in all tested cancer cell lines compared to those in HEK 293A. Remarkably, hsa-mir-1229-3p is upregulated in cell lines derived from metastatic sites and downregulated in primary tumor lines. However, further studies are required to assess whether miR-1229-3p is associated with the metastatic potential of human tumors.
To further increase the predictive value for clinical applications, we extended studies of these mirtronic miRNAs to a number of primary pancreatic, colorectal, stomach cancerous and healthy tissues (Fig. 3). We presume that analysis of non-paired cohorts of tumoral and non-tumoral samples allows a better estimation of miRNA expression changes, since healthy tissues are certainly not affected by cancerogenesis. Three mirtron-derived miRNAs showed organ-specific profiles in cancerous tissues of the digestive system. T3 stage pancreatic tumors exhibited a decline of the hsa-miR-1227-3p and hsa-miR-1236-3p expression. The levels of hsa-miR-1226-3p decreased in the T3 stage colorectal but conversely increased in the stomach cancer tissues. Remarkably, hsa-mir-1229-3p, which was upregulated in the metastatic site-derived cell lines, exhibited no expression change in the primary tumors. Previous studies demonstrated that cellular levels of hsa-miR-1226-3p and hsa-miR-1236-3p inversely correlate with cancerogenesis in breast and hepatoma cancer cells, respectively [20, 33]. Taken together, these results suggest that dysregulation of conventional mirtronic miRNAs could lead to the initiation and progression of specific human cancers. Moreover, the development of new medical applications can be based on artificial mirtron-mediated gene silencing platforms. As promising and affordable tools, they surpass conventional RNA interference (RNAi) approaches and allow to consolidate the high therapeutic potential of RNAi and protein-coding genes in specific cell types [34–36]. For example, hsa-miR-1226-3p processed from an artificial conventional mirtron effectively silences aberrant myotonic dystrophy protein kinase , mmu-miR-1224 leucine-rich repeat serine/threonine-protein kinase 2, and α-synuclein Parkinson disease-associated genes , while an artificial 3′-tailed mirtron knocks down the expression of the vascular endothelial growth factor A .
It is known from previous studies that canonical miRNA biogenesis and splicing mechanisms are directly coupled . The binding of splicing regulatory proteins SRSF1 and hnRNPA1 to the stem loop of pri-miR-7 and pri-miR-18a, respectively, appears to affect Drosha processing of primary miRNA transcripts [26, 33]. Similarly, KHSPR promotes the maturation of a subset of miRNA precursors, such as pre-let-7a, pre-mir-1, and pre-mir-15 . Consequently, SFs regulate both splicing and canonical miRNA processing, but little is known about their function in the biogenesis of miRNAs hosted on mirtrons. It has been only observed in patients with myelodysplastic syndrome that SF3B1 and SRFS2 mutations are associated with a downregulated expression of miRNAs derived from the putative 5′-tailed mirtrons hsa-miR-3605-5p and hsa-miR-4728-5p, respectively .
Our results revealed that increased nuclear levels of SRSF1 in tested HCT116 cells significantly enhanced the hsa-miR-1229-3p expression, while upregulated SRSF2 increased the abundance of the mirtronic miRNAs hsa-miR-1227-3p and hsa-miR-1229-3p. To our knowledge, these data, for the first time, demonstrate that an individual SF can directly act as a positive regulator for particular species of mirtronic miRNAs.
To gain insight into the regulation of mirtronic miRNA by components of splicing machinery, we correlated the mirtronic miRNA levels with the expression profiles of five SFs in different human digestive and excretory system cell lines or pancreatic, colorectal, and stomach tissues. It should be noted that RNA (this paper) and proteins  were purified simultaneously from the same tissue or cell line samples. The experiments revealed a reduced expression of SRSF1 and SRSF2 in pancreatic, kidney, colorectal, and stomach cancer cell lines as well as in cancer tissues — SRSF2 was downregulated in stomach tumors, while SRSF1 was reduced in all types of tested tumors . Accordingly, tumor-promoting roles of SRSF1 and SRSF2 have been reported in various cancer types [40, 41]. Still, reduced expression of SRSF1 and SRSF2 proteins in the cancer cells  weakly correlated with signature of mirtronic miRNA expression (Tables 2 and 3). For example, no obvious differences in abundance of hsa-miR-1229-3p were seen among samples collected from the pancreas, colon, or stomach, notwithstanding a ~14-fold under-expression of SRSF1 and a sevenfold of SRSF2 in the colorectal and stomach cancerous tissues. At a first glance, these results seem to contradict the finding that these SFs positively modulate the levels of mirtronic miRNAs under overexpression conditions in the model cell line HCT116 (Fig. 4). However, first, a dual role for SRSF1 as a splicing activator and repressor was proposed . Second, the expression of other factors involved in the regulation of splicing, U2AF35, U2AF65, and KHSRP, also significantly varied among cell lines and tumors . In addition, the spliceosome mutations may affect the expression of miRNA genes . Thus, biogenesis of mirtronic miRNAs could be regulated by different SFs or by a network of antagonistic and collaborative interactions of spliceosome components. In general, our data imply that a combination of multiple SFs may differently affect the biogenesis of mirtron-derived miRNAs, making it extremely difficult to predict particular effects of individual spliceosome components on the mirtron processing.
In this study, we experimentally validated three novel splicing-dependent miRNAs processed from mirtrons, hsa-miR-1227-3p, hsa-miR-1229-3p, and hsa-miR-1236-3p. The expression analysis, for the first time, identified specific profiles of mirtronic miRNAs in cancerous cells and tumors suggesting their potential for exploiting as diagnostic tools. Finally, we demonstrated that the SRSF1 and SRSF2 SFs contribute to the biogenesis of mirtronic hsa-miR-1227-3p and hsa-miR-1229-3p.
Human cell cultures
HEK 293A (embryonic kidney), PANC-1 (pancreas/duct, epithelioid carcinoma), SU.86.86 (pancreas, ductal carcinoma; derived from a metastatic site: liver), T3M4 (pancreas, ductal carcinoma; metastasis), CaKi-1 (kidney, clear cell carcinoma, derived from a metastatic site: skin), 786-O (kidney, renal cell adenocarcinoma), HCT116 (colon, colorectal carcinoma), and KATOIII (stomach, gastric carcinoma; derived from a metastatic site: pleural effusion and supraclavicular and axillary lymph nodes and Douglas cul-de-sac) cells were cultured in the RPMI 1640 medium supplemented with 10 % fetal bovine serum and penicillin (100 U/ml) and streptomycin (100 μg/ml).
Tumors and healthy tissues
Collection of surgically removed tumor and healthy tissue samples was described previously . In total 29 healthy (8 pancreatic, 6 colorectal, 15 stomach) ant 37 tumor (12 pancreatic, 12 colorectal, 13 stomach) tissues were used for the experiments. The experiments were undertaken with the understanding and written consent of each subject, and the study conforms to The Code of Ethics of the World Medical Association (Declaration of Helsinki).
Construction of plasmids
Minigene constructs of 1226/DHX30 (from exon 19 to exon 21; mirtronic intron 20), 1227/PLEKHJ1 (exons 5 to 7; mirtronic intron 5), 1229/MGAT4B (exons 11 to 13; mirtronic intron 12), 1236/NELFE (exons 3 to 4; mirtronic intron 3), 1238/ATG4D (exons 7 to 9; mirtronic intron 8), 3064/DDX5 (exons 10 to 12, mirtronic intron 11), 3940/KHSRP (exons 13 to 15, mirtronic intron 14), 6515/CARL (exons 5 to 7, mirtronic intron 5), and 6850/RPL8 (exons 2 to 4, mirtronic intron 2) were amplified from human Jurkat cell genomic DNA (ThermoFisher Scientific) using Phusion High-Fidelity DNA Polymerase (ThermoFisher Scientific) according to the manufacturer’s instructions. PCR products were digested with BamHI and HindIII restriction endonucleases, except 3940/KHSRP, 6515/CARL (digested with HindIII and EcoRI), and 6850/RLP8 (EcoRI and NotI), and inserted into pcDNA3 plasmid. PCR introduced 5′- and 3′-splice site mutations by using Phusion High-Fidelity DNA Polymerase with primers containing splicing site mutations (Additional file 2: Table S2). Amplification products were circularized using T4 DNA ligase (all enzymes were obtained from ThermoFisher Scientific). All final plasmid constructs were confirmed by Sanger sequencing.
The expression plasmids pSRSF1 and pSRSF2 were constructed by insertion of SRSF1 isoform 1 and SRSF2-coding DNA sequence into the pcDNA3 vector, respectively. Total RNA, extracted from HCT116 cells, was converted to complementary DNA (cDNA) using RevertAid RT kit (ThermoFisher Scientific). The CDS of genes were PCR-amplified using primers containing HindIII and BamHI sites (Additional file 2: Table S1), digested with appropriate restriction enzymes and the resulting fragments ligated into pcDNA3. Clones were confirmed by DNA sequence analysis.
HTC116 cells were cultured in RPMI 1640 media. Transient transfections were performed using Lipofectamine LTX transfection reagent (Invitrogen) according to the manufacturer’s instructions. For overexpression assay, HTC116 cells were seeded in 60-mm plates and allowed to grow for 24 h prior to transfection with pcDNA3-SRSF1, pcDNA3-SRSF2, pcDNA3-DHX30, pcDNA3-PLEKHJ1, pcDNA3-MGAT4B, pcDNA3-NELFE, pcDNA3-ATG4D, pcDNA3-DDX5, pcDNA3-KHSRP, pcDNA3-CARL, or pcDNA3-RPL8 constructs. A pcDNA3 vector was transfected to be used as a RT-qPCR reference. RNAs and proteins were isolated after 48 h. The experiments were independently repeated three times.
RNA from cell lines was isolated using an RNAzol® RT reagent according to the manufacturer’s instructions (Molecular Research Center). Native proteins for SF studies by Jakubauskienė et al.  and total RNA used for the miRNA analysis (present paper) from tumors and healthy tissues were purified from the same sample using mirVana™ PARIS™ kit (ThermoFisher Scientific) according to the manufacturer’s instructions. Fraction of short RNAs (<200 nt) was used for miRNA RT and qPCR experiments. The integrity of long RNAs (>200 nt) was assessed using an Agilent 2100 Bioanalyzer or agarose (1 %) gel electrophoresis. To avoid DNA contamination before RT reaction, the RNA samples were treated with DNase I (ThermoFisher Scientific) according to the manufacturer’s instructions.
Real-time RT-PCR of miRNAs
Each 20 μl of the RT reaction in the RT buffer contained 200 units of RevertAid Reverse Transcriptase (ThermoFisher Scientific), 1 mM dNTP (ThermoFisher Scientific), 1 μM RT primer mixture (Metabion), 16 units of RNase inhibitor RiboLock (ThermoFisher Scientific), and 200 ng of RNA. The mixture was incubated for 20 min at 25 °C and for 60 min at 37 °C and then heat-inactivated for 10 min at 70 °C. Real-time PCR was performed with SYBR Green PCR Master Mix (ThermoFisher Scientific) (35 cycles): initial denaturation for 10 min at 95 °C, followed by three cycles of amplification 15 s at 95 °C, 1 min at 55 °C, and 30 s at 60 °C, then 32 cycles 10 s at 95 °C and 30 s at 60 °C. Relative quantification of changes in the miRNA expression levels was performed using the comparative C t (threshold cycle) method with normalization to the expression of endogenous control RNU48. Specific miRNA and RNU48 primers are provided in Additional file 2: Table S3. The RT primer mixture was composed of two primers: specific miRNA and RNU48. Quantitative PCR analysis was carried out on Rotor-Gene 6000 (Corbett Life Science) equipment. Part of RT-PCR products were inserted into pUC19 plasmid, and constructs were confirmed by sequence analysis.
Detection of spliced and unspliced forms of mRNA
Spliced and unspliced DHX30, PLEKHJ1, MGAT4B, NELFE, ATG4D, DDX5, KHSRP, CARL, and RPL8 mRNA isoforms were detected by RT-PCR. cDNA synthesis was carried out using RevertAid Reverse Transcriptase and random hexamer primers according to the manufacturer’s (ThermoFisher Scientific) instructions. PCR was carried out using TrueStart Hot Start Taq DNA Polymerase (ThermoFisher Scientific) according to the manufacturer’s instructions. Amplification products were separated on 1 % agarose gel. TBP was used as loading control. Sequences of primers specific for the outer exons of mirtronic introns are provided in Additional file 2: Table S4.
Protein preparation from cell cultures and western blotting
The nuclear fractions of proteins for western blotting were prepared from cell cultures using the NE-PER Nuclear and Cytoplasmic Extraction Reagents kit (ThermoFisher Scientific) according to the manufacturer’s instructions. Proteins were separated on a 10 % SDS–polyacrylamide gel and transferred onto the membrane. The membrane was incubated with appropriate primary antibody: anti-SRSF1 (Santa Cruz), anti-SRSF2 (Abcam), or anti-β-actin (Abcam), and washed and incubated with secondary antibody (Dako). The membrane was developed using TMB reagent (Sigma-Aldrich). All data were quantitated using MultiGauge software (Fujifilm). β-Actin expression was used for data normalization.
Statistical analysis was performed using Sigma Plot software v. 11. Two-tailed unpaired Student’s t test and Mann-Whitney rank sum test were used to compare the differences in distribution between experimental results. A value of p < 0.05 was considered statistically significant.
We thank Antanas Gulbinas for the helpful discussion and Alexandra Plotnikova and Saulius Klimašauskas for the critical comments on the manuscript. This work was funded by the Research Council of Lithuania grant LIG-10/2012.
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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