A novel regulatory network among LncRpa, CircRar1, MiR-671 and apoptotic genes promotes lead-induced neuronal cell apoptosis
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Lead is a metal that has toxic effects on the developing nervous system. However, the mechanisms underlying lead-induced neurotoxicity are not well understood. Non-coding RNAs (ncRNAs) play an important role in epigenetic regulation, but few studies have examined the function of ncRNAs in lead-induced neurotoxicity. We addressed this in the present study by evaluating the functions of a long non-coding RNA (named lncRpa) and a circular RNA (named circRar1) in a mouse model of lead-induced neurotoxicity. High-throughput RNA sequencing showed that both lncRpa and circRar1 promoted neuronal apoptosis. We also found that lncRpa and circRar1 induced the upregulation of apoptosis-associated factors caspase8 and p38 at the mRNA and protein levels via modulation of their common target microRNA miR-671. This is the first report of a regulatory interaction among a lncRNA, circRNA, and miRNA mediating neuronal apoptosis in response to lead toxicity.
KeywordsCircRNA LncRNA MiRNA Cell apoptosis Lead Neurotoxicity
Lead is an industrial and environmental pollutant that can cause pathological changes to multiple organ systems, including the nervous system. Lead has irreversible neurotoxic effects on the developing brain (Wang et al. 2008). However, the molecular mechanisms of lead-induced neurotoxicity remain unclear. There have been few studies investigating the role of non-coding RNAs (ncRNAs) in this process, although they have been shown to play important roles in many biological processes, including transcriptional regulation, DNA replication, RNA processing, mRNA stability and translation, and protein degradation and transport (Storz 2002).
Long non-coding RNAs (lncRNAs) and circular RNAs (circRNAs) have been the focus of many recent studies on ncRNA function. LncRNAs are 200 nucleotides in length and are distributed in the nucleus or cytoplasm (Maruyama and Suzuki 2012; Okazaki et al. 2002). LncRNAs regulate gene expression at the transcriptional, post-transcriptional, and epigenetic levels and have been implicated in species evolution, embryonic development, metabolism and disease (Mercer et al. 2009; Taft et al. 2010). On the other hand, knowledge of circRNA functions is limited. These molecules were originally identified in an RNA virus (Wilusz and Sharp 2013); they are widely present in mammalian cells and have been linked to the regulation of gene expression (Zhang et al. 2014), alternative splicing (Lasda and Parker 2014), and translation through interaction with RNA-binding proteins (Zhang et al. 2013). CircRNAs have been shown to bind microRNAs (miRNAs) as sponges, thereby indirectly regulating target gene expression (Hansen et al. 2013a; Wilusz and Sharp 2013). There have been no reports to date of interactions between lncRNAs and circRNAs.
The present study investigated the functions of lncRNAs and circRNAs in lead-induced neurotoxicity. We first carried out an RNA screen in a mouse model. A quantitative real-time polymerase chain reaction (qRT-PCR) analysis revealed that the pro-apoptotic lncRNA (named lncRpa) and the apoptosis-related circRNA (named circRar1) were upregulated in the hippocampus and cerebral cortex of mice with lead-induced neurotoxicity. A similar upregulation of lncRpa and circRar1 was observed in N2a cells treated with lead acetate (PbAc). We found that lncRpa and circRar1 acted via the common target miR-671 to promote neuronal apoptosis. These findings highlight the regulatory roles of lncRNAs and circRNAs in lead-induced neurotoxicity and provide the first evidence of these ncRNAs modulating a cellular process by jointly targeting a specific miRNA.
Materials and methods
Lead-induced neurotoxicity models
The mouse model of lead-induced neurotoxicity has been previously described, and the animal studies were approved by the Animal Care and Use Committee of Guangzhou Medical University (Nan et al. 2016). We also used mouse neuroblastoma N2a cells exposed to PbAc at a concentration of 0.1 µM for 48 h as an in vitro model of lead-induced neurotoxicity (Nan et al. 2016).
RNA was obtained from brain tissue (cerebellum, pons, medulla oblongata, hippocampus, and cerebral cortex) of mice with lead-induced neurotoxicity (2 and 5 weeks of exposure to PbAc) and control mice. Trizol reagent (Invitrogen, Carlsbad, CA, USA) was used according to the manufacturer’s instructions to extract total RNA from tissues and cells. For quantitation of circRNAs, RNase R (Epicentre, Madison, WI, USA) was added to degrade linear RNAs. RNA quality and concentration were measured with a NanoDrop1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE, USA).
High-throughput RNA sequencing
The HiSeq 2000 sequencing platform (Illumina, San Diego, CA, USA) was used for high-throughput RNA sequencing. The protocol involved removal of rRNA, followed by synthesis of double-stranded cDNA and end repair. After linking sequencing adaptors and selecting fragments, the second strand of cDNA was degraded and the remaining strand was enriched by PCR. The quality of the library was confirmed by sequencing. A bioinformatic analysis of the raw sequencing data was carried out. Differentially expressed ncRNAs were searched in the NCBI database (http://www.ncbi.nlm.nih.gov/) to determine their genome loci.
The Goscript Reverse Transcription System (Promega, Madison, WI, USA) was used to reverse transcribe lncRNAs, circRNAs, and mRNAs to cDNA. Go Taq qPCR Master Mix (Promega) was used for qRT-PCR. All-in-one miRNA qRT-PCR Detection kit (Genecopoeia, Rockville, MD, USA) was used to reverse transcribe and amplify miRNAs. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as an internal control for the relative quantitation of lncRNAs, circRNAs, and mRNAs, whereas U6 was used for miRNAs. The detection of internal control gene GAPDH would be affected after treating with RNase R; we divided the same RNA sample into two uniform parts when performing the qRT-PCR experiment. One part was treated with RNase R for delinearization; this part was for the further detection of circRNA. The other part was treated with RNase R-free water for finally detecting GAPDH gene. The primer sequences are shown in Supplementary Table 3. The 2−ΔΔCt method was used to determine relative expression levels.
RNA interference and overexpression
LncRNA and circRNA expression was suppressed by siRNA-mediated knockdown. Three different siRNAs were designed and tested for both lncRpa and circRar1. Overexpression vectors for lncRpa and circRar1 were also constructed (BersinBio, Guangzhou, China). CircRNA upstream intron cyclization component (526 bp), circRNA (462 bp) and circRNA downstream intron cyclization component (804 bp) were included in circRNA expression area. BamHIand Hind III were jointly connected to expression vector pcDNA 3.1+ through double enzyme connection. Overexpression and siRNA sequences are shown in Supplementary Table 1. A specific inhibitor and mimic (RiboBio, Guangzhou, China) were used to inhibit or induce miR-671 expression, respectively. Cells were transfected with plasmid using EndoFectin Lenti reagent (Genecopoeia). RiboFECT CP Transfection kit (166T) (RiboBio) was used for the miR-671 inhibitor and mimic.
Detection of cell apoptosis by FCM
The Annexin V-fluorescein isothiocyanate (FITC) apoptosis assay kit (KeyGen Biotech, Nanjing, China) was used according to the manufacturer’s instructions to detect apoptotic cells 48 h after transfection and PbAc treatment. Briefly, 5 × 105 cells were collected and resuspended in 100 μl 1 × binding buffer. Five microliters Annexin V-FITC and 5 μl propidium iodide staining solution were added to the cells, followed by incubation at room temperature (shielded from light) for 10 min. Four hundred microliters 1 × binding buffer was added to the reaction, and cells were analyzed by FCM (BD Biosciences, Franklin Lakes, NJ, USA) within 1 h.
Detection of cell apoptosis by TUNEL assay
A TUNEL kit (Roche Diagnostics, Indianapolis, IN, USA) was used to detect apoptotic cells. Cells were cultured on Lab-Tek chambered slides (Thermo Fisher Scientific, Waltham, MA, USA). Following treatment, the samples were washed twice with phosphate-buffered saline (PBS) and fixed with 4 % paraformaldehyde at room temperature for 20 min, followed by two washes with PBS. Proteinase K (20 μg/ml; Sangon Biotech, Shanghai, China) was added, and the slides were covered with a film and incubated at 37 °C for 20 min and then washed twice with PBS. The TUNEL reaction mixture (enzyme and labeling solutions at a 1:9 ratio) was added to the slides, which were covered with film and incubated at 37 °C for 60 min. After three washes with PBS, converter-peroxidase was added at 37 °C for 30 min; after three more washes with PBS, diaminobenzidine reagent (Roche Diagnostics) was added at room temperature for 10 min. The samples were washed three times with PBS and counterstained with hematoxylin for 10 s and then washed with running water. After dehydration in a graded series of alcohol, the samples were dried and mounted with neutral balsam. Nuclei with yellowish brown staining were positive (apoptotic), and hematoxylin-counterstained intact nuclei appeared blue under a light microscope.
CCK-8 cell viability assay
The CCK-8 assay kit (Beyotime Institute of Biotechnology, Shanghai, China) was used to assess cell viability. Cells were harvested in logarithmic phase, and 100 μl of the suspension (~2000 cells) were seeded in each well of a 96-well plate and incubated overnight at 37 °C and 5 % CO2. EndoFectin Lenti reagent, plasmids, siRNAs, and serum-free Dulbecco’s Modified Eagle’s Medium (DMEM; HyClone, Logan, UT, USA) equilibrated to 15–25 °C were added to the wells followed by incubation at room temperature for 10–25 min. The medium was changed after 6 h, and 0.1 μM PbAc solution was added for 48 h. Ten microliters CCK-8 solution was added for 1–4 h, and the absorbance at 450 nm was measured using a microplate reader.
Total protein was extracted using a commercial kit (KeyGen Biotech). Protein samples (4–8 μg/μl) were mixed with a 4:1 ratio of 5 × loading buffer and β-mercaptoethanol and stored at −80 °C until use. Proteins (40–60 μg per well) were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (100–120 V). A protein marker with a molecular weight range of 16–220 kDa was used as reference. The proteins were transferred to a polyvinylidene difluoride membrane (Merck Millipore, Billerica, MA, USA) at 200 mA using a wet membrane-transfer device (Bio-Rad, Hercules, CA, USA). The membrane was washed with Tris-buffered saline containing 0.1 % Tween-20 (TBST) for 1–2 min and then blocked at room temperature for 60 min with TBST containing 5 % non-fat milk powder. After overnight incubation at 4 °C with primary antibodies, the membrane was washed with TBST three times for 15 min. The membrane was then incubated at room temperature for 60 min with secondary antibody and washed three times with TBST for 15 min each. Protein bands were visualized using the BeyoECLPlus chemiluminescence reagent (Beyotime Institute of Biotechnology) followed by exposure to X-ray film. Primary antibodies against the following proteins were used in this study: caspase3 (Cell Signaling Technology, Danvers, MA, USA), caspase9 (Epitomics, Burlingame, CA, USA), Akt2 (Cell Signaling Technology), caspase 8 (Proteintech, Rosemont, IL, USA), and p38 (Cell Signaling Technology). The secondary antibody was horseradish peroxidase-conjugated IgG (Boster Bio, Pleasanton, CA, USA).
Cells grown on coverslips were fixed with 4 % paraformaldehyde at room temperature for 15 min, washed twice with 0.1 % diethylpyrocarbonate solution and treated with 0.5 % Triton X-100 at room temperature for 5 min. The samples were dehydrated in a graded series of alcohol and air-dried. After adding probe hybridization solution, the samples were mounted, denatured at 73 °C for 3 min, and hybridized in a humid and dark environment at 37 °C for 12–16 h with Cy3-labeled miRNA probe, 6-carboxyfluorescein-labeled circRNA probe, and Cy5-labeled lncRNA probe (BersinBio). The samples were washed three times with a pre-heated (43 °C) solution consisting of 50 % formamide and 2× saline sodium citrate (SSC), and then washed twice with 2× SSC (37 °C). After counterstaining with 4′, 6-diamidino-2-phenylindole, the samples were mounted with fluorescence mounting medium and imaged with a microscope.
Dual luciferase reporter gene assay
Cells were seeded and incubated for 24 h. At 80–90 % confluence, the cells were transfected with firefly and Renilla luciferase plasmids. After washing with PBS, passive lysis buffer (PLB) was added and cells were incubated at room temperature for 15 min, with a micro-oscillator used to lyse the cells. Lysates were centrifuged at 10,000 rpm for 5 min at 4 °C. The supernatants were removed, and 20 μl sample were transferred to a 96-well plate and mixed with 100 μl Dual-Glo Luciferase Assay System (Promega), with cell lysis buffer used as the control. The relative light units were measured before and after adding 100 μl Stop & Glo reagent.
RNA antisense purification (RAP)
The RAP kit (BersinBio) was used for this experiment. RAP employs specific biotinylated probes that hybridize to target RNAs (mRNAs or miRNAs); these can then be pulled down, reverse transcribed to cDNA, and identified by qRT-PCR or sequencing. A total of 107 cells were washed with PBS and cross-linked by ultraviolet irradiation at 254 nm (0.15 J cm−2). Cells were lysed with 1 ml lysis buffer and fully homogenized with a 0.4 mm syringe. Two different 45-bp biotinylated antisense probes (0.2 nmol) were added to the lncRNA-RAP system and one 50-bp biotinylated antisense probe (0.2 nmol) targeting the adaptor sequence was added to the circRNA-RAP system. The probes were denatured at 65 °C for 10 min and hybrided at room temperature for 2 h before adding 200 µl streptavidin-coated magnetic beads. Non-specifically bound RNAs were removed by washing, and Trizol reagent was used to recover miRNAs specifically interacting with ncRNAs. PCR and qRT-PCR were used to analyze binding strength after reverse transcribing the miRNAs. The probes are shown in Supplementary Table 4.
Data are presented as mean ± SD. All experiments were performed at least three times, and western blotting, TUNEL, FCM, and FISH results are representative of three independent experiments. The unpaired t test was used for statistical analyses. * represents statistically significant difference (p < 0.05). ** represents highly statistically significant difference (p < 0.01). Data were analyzed using SPSS v.19.0 software (IBM, Armonk, NY, USA).
Identification of lncRNAs and circRNAs differentially expressed in lead-induced neurotoxicity
LncRpa and circRar1 promote apoptosis in lead-induced neurotoxicity
LncRpa and circRar1 interact directly with miR-671
LncRpa and circRar1 regulate miR-671 expression
MiR-671 inhibits apoptosis
MiR-671 regulates apoptosis-associated factors
LncRpa and circRar1 regulate apoptosis-associated factors in lead-induced neurotoxicity
High-throughput sequencing technology has broadened our understanding of gene regulatory networks. Whole genome sequencing has revealed that about 93 % of the genome is transcribed as RNA, but only 2 % encode proteins (Birney et al. 2007). Although the total number of nucleotides in the human genome is 30 times that of the nematode genome, the number of protein-coding sequences is comparable, which highlights the importance of ncRNA sequences in the regulation of eukaryotic gene expression (Costa 2008).
Environmental toxins such as lead can adversely affect human health, but the molecular mechanisms of lead-induced neurotoxicity are not well understood. Most research in this area has focused on the role of mRNAs (Soliman et al. 2015; Gao et al. 2016) or miRNAs (Li et al. 2015; Martinez-Pacheco et al. 2014), and there is little, if any information on how lncRNAs and circRNAs are involved in lead-induced neurotoxicity. The roles of ncRNAs have been extensively investigated in the context of carcinogenesis and cancer development (Cheng et al. 2015). For example, H19 is aberrantly expressed in many types of cancer such as liver and bladder cancers and pancreatic ductal carcinoma (Ma et al. 2014; Luo et al. 2013; Tsang and Kwok 2007), while HOTAIR has been implicated in various aspects of cancer development (Wu et al. 2014). Genome-wide association studies have shown that most cancer risk loci are found in non-coding sequences (Cheetham et al. 2013). Less is known about circRNAs, despite their prevalence in mammalian cells. These molecules also regulate gene expression at the post-transcriptional level (Memczak et al. 2013), and some have been found to be associated with tumors (Hansen et al. 2013b; Peng et al. 2015; Zhao and Shen 2015).
NcRNAs interact in a complex regulatory network (Supplementary Fig. 1). LncRNAs alter chromatin structure via activation and transport of relevant proteins (Zhao et al. 2008; Tsai et al. 2010; Yao et al. 2010) and are also involved in the regulation of transcription factors (Hung et al. 2011) and mRNA and protein expression (Gong and Maquat 2011; Yoon et al. 2012). In many instances, lncRNAs carry out their functions by modulating the expression of miRNAs at the level of transcription, post-transcription, or splicing (Poliseno et al. 2010; Augoff et al. 2012; Steck et al. 2012; Wang et al. 2010). MiRNAs play a critical role in this regulatory network by directly targeting mRNAs (Orom et al. 2008). Previous studies have shown that miRNAs can be adsorbed by circRNAs, which act as endogenous miRNA competitors (Hansen et al. 2013a, b; Xu et al. 2015).
In this study, we found that lncRNA lncRpa and circRNA circRar1 were differentially expressed in lead-induced neurotoxicity and directly regulated miR-671 expression to promote neuronal apoptosis via upregulation of the pro-apoptotic proteins caspase8 and p38. We also found that miR-671 negatively regulate circRar1 and lncRpa. Thus, lncRpa and circRar1 jointly target miR-671 to modulate the expression of apoptosis-associated proteins in lead-induced neuronal apoptosis. These findings highlight a new mechanism of lead-induced neurotoxicity and provide a insight for the future investigations of the pathological process.
We thank Dr. Sijin Liu (professor of Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences) for recommendations on the manuscript. This work was supported by the National Key Basic Research Program of China (No. 2012CB525004 to Y. G. J.), the National Natural Science Foundation of China (No. 21277036, 81573180, and 81172633 to Y. G. J.), the Key Program of Guangdong Natural Science Foundation (No. 2014A030311017 to Y. G. J.), the University Major Program of Guangdong (No. 2014SZD48 to Y. G. J.), the University Talent Program of Guangdong (No. 2013-164 to Y. G. J.), and the University Chief Scientist Program of Guangzhou (No. 1201541575 to Y. G. J.).
Y. G. J. obtained funding for the study. Y. G. J., A. R. N., C. F. Y., and Z. S. W. designed the experiments. L. J. C. and N. Z. performed qRT-PCR and analyzed the data. Z. Z. L. and T. Y. carried out the Luciferase Reporter Gene Assay and analyzed the data. A. R. N. performed all other experiments and analyzed the data. A. R. N. and Y. G. J. wrote the manuscript. All authors read and approved the final manuscript.
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