miR-15b/16-2 Regulates Factors That Promote p53 Phosphorylation and Augments the DNA Damage Response following Radiation in the Lung

Background: MicroRNAs play an integral role in modulating cellular stress, including DNA damage.

Results: miR-15b induces p53 phosphorylation, DNA repair, apoptosis, and cell cycle arrest while inhibiting Wip1 expression following irradiation in lung epithelial cells.

Conclusion: The miR-15b/Wip1 axis could be a potential therapeutic target in radiation-associated lung disease.

Significance: MicroRNAs may represent an actionable target in the lung following radiation exposure.

Abstract

MicroRNAs (miRNAs) are regulatory RNAs frequently dysregulated in disease and following cellular stress. Investigators have described changes in miR-15b expression following exposure to several stress-inducing anticancer agents, including ionizing radiation (IR), etoposide, and hydrogen peroxide. However, the role for miR-15b as a mediator of cellular injury in organs such as the lung has yet to be explored. In this study, we examined miR-15b expression patterns as well as its potential role in DNA damage and repair in the setting of IR exposure. We showed that miR-15b is up-regulated in a dose- and time-dependent manner in human bronchial epithelial cells following IR. miR-15b expression was highest after 2 h of IR and decreased gradually. Survival rates following IR were also higher in miR-15b/16-2-overexpressing cells. Cell cycle arrest in G₂/M phase and an increased DNA repair response were observed in IR-exposed miR-15b/16-2 stable cells. We observed an up-regulation of components of the ataxia telangiectasia mutated (ATM)/Chek1/p53 pathway in miR-15b/16-2-overexpressing cells after IR. Moreover, a pathway-based PCR expression array of genes demonstrated that miR-15b/16-2 overexpression significantly induced the expression of genes involved in ATM/ataxia telangiectasia and Rad-3-related (ATR) signaling, apoptosis, the cell cycle, and DNA repair pathways. Here we demonstrated a novel biological link between miR-15b and DNA damage and cellular protection in lung cells. We identified Wip1 (PPM1D) as a functional target for miR-15b and determined that miR-15b induction of the DNA damage response is partially dependent upon suppression of Wip1. Our study suggests that miR-15b/Wip1 could be a potential therapeutic target in radiation-induced lung disease.

Introduction

MicroRNAs (miRNAs)² are small RNA molecules ∼22 nucleotides in length that have recently emerged as critical regulators of gene expression (1). The importance of miRNAs in cell cycle progression, cell proliferation, genomic instability, and malignant transformation is increasingly being recognized (2–4). The miR-15a/16-1 and miR-15b/16-2 clusters have been shown to regulate cell cycle and apoptosis by targeting Cyclin D3 (CCND3), Cyclin E1 (CCNE1), CDK6 (5, 6), and the antiapoptotic gene Bcl-2 in various cancer cells (7). Investigators have demonstrated that the miR-15b/16-2 cluster is an important regulator of gene expression that fine-tunes complicated gene networks in a variety of cell lines. Therapeutic strategies targeting multidrug resistance-related miRNAs, such as miR-15b and miR-16, may be more promising than targeting single proteins, considering the fact that miRNAs function through the regulation of multiple genes, leading to a broader impact on diverse cellular processes. miR-15b regulates cell cycle progression in glioma cells by targeting cell cycle-related molecules (8). Overexpression of miR-15b resulted in cell cycle arrest at G₀/G₁ phase, whereas suppression of miR-15b expression resulted in a decrease in the number of cells in G₀/G₁ phase and a corresponding increase in cells in S phase. miR-15b is up-regulated in both head and neck squamous cell carcinoma and oral squamous cell carcinoma but, following hypoxic conditions, is down-regulated in nasopharyngeal carcinoma (9). The pathophysiological functions of miR-15b have yet to be thoroughly investigated. Recently, it has been reported that the expression of miR-15b may be altered following exposure to various genotoxic stressors, including radiation, hydrogen peroxide (H₂O₂), and etoposide (10). However, whether or not miR-15b mediates cellular stress and damage is unknown. Here we examined the expression patterns and potential role for miR-15b in DNA damage and repair in human bronchial epithelial cells.

Cellular stress targets numerous biological processes in human cells, including cell cycle progression, gene transcription and translation, as well as epigenetic mechanisms such as DNA damage and repair. The DNA damage response (DDR) is a signaling network initiated by lesion recognition and amplified by multiple mediator signaling proteins that eventually activate downstream effectors to modulate cell fate by manipulating gene expression, cell cycle control, DNA repair, apoptosis, and senescence (11). Our results suggest that miR-15b/16-2 plays an important role in DDR. miR-15b/16-2 overexpression induced G₂/M phase arrest following IR in bronchial epithelial cells. Multiple DNA repair pathways have evolved to resolve various DNA lesions. These pathways include single strand break repair, homologous recombination, non-homologous end joining, nucleotide excision repair, mismatch repair, translation synthesis, the Fanconi anemia pathway, and others. Recently, studies have suggested that alterations in miRNA levels resulting from either mutations or aberrant expression correlate with various human cancers (12). Furthermore, abnormally expressed miRNAs in human cancers target transcripts of essential protein-coding genes involved in tumorigenesis, including oncogenes and tumor suppressor genes (13).

It has been reported that γ-H2AX is phosphorylated by ATM to further recruit partners such as BRCA1 and TP53BP1 and form a scaffold around the DNA break site (14). The DNA damage signal is then transduced by ATM/ATR kinase to downstream pathways such as the cell cycle, DNA repair, or apoptosis. It has recently been reported that ATM expression is also under the control of miRNAs (15). miR-421 and miR-101 are reported to suppress ATM expression by targeting the 3′UTR of ATM transcripts.

In this study, we showed that irradiation increased the expression of miR-15b shortly after exposure, with a peak at 2 h followed by a gradual decrease in expression. Following IR exposure, 15b/16-2 stable cells exhibited cell cycle arrest at G₂/M phase, increased DNA damage and repair, and increased cell survival. We also found that miR-15b expression correlated inversely with Wip1 (PPM1D) expression following IR. It has been reported recently that Wip1 functions as a homeostatic regulator of the ATM-mediated signaling pathway in response to IR (16). Wip1 is a serine/threonine phosphatase that has been shown to inactivate the DDR through several mechanisms, including dephosphorylation of p53, γ-H2AX, and ATM. It has also been demonstrated that down-regulation of Wip1 is required for maintenance of permanent G₂ arrest (17). Wip1 is a novel target of miR-15b, and deregulation of the miR-15b/Wip1 may be a mechanism for regulating the DDR.

EXPERIMENTAL PROCEDURES

Cell Culture, Transfection, and Irradiation

A human bronchial epithelial cell (HBEC) line was obtained from the ATCC. EV (empty vector) and miR-15b/16-2 stable cells were modified from HBECs. HBECs were infected with the human pre-miR expression construct lentiviral-miR-15b/16-2 (LV-miR-15b/16-2) (catalog no. PM1RH15BPA, SBI) or GFP lentiviral empty vector (LV-EV) according to the instructions of the manufacturer. Briefly, HBECs were infected with LV-miR-15b/16-2 or LV-EV in the presence of 4 μg/ml Polybrene (Sigma-Aldrich, St. Louis, MO) at a multiplicity of infection of 10 and centrifuged at 1800 rpm for 5 min. Thereafter, cells were cultured for 48 h and analyzed by fluorescence-activated cell sorting (BD Biosciences). The expression of mature miR-15b and miR-16-2 was confirmed by quantitative real-time PCR (qRT-PCR). Cells were maintained in keratinocyte-serum-free medium (Invitrogen) supplemented with human recombinant EGF, bovine pituitary extract, and 1% penicillin/streptomycin (Invitrogen). All transfections were performed using FuGENE6 according to the recommendations of the manufacturer (Roche). To knock down miR-15b, HBECs were transfected with silencer-miR-15b and scramble miRNA (Applied Biosystem) for 24 h. Irradiation was performed using a RS-2000 biological irradiator (Rad-Source) with 160-kV x-rays with a 0.3-mm copper filter in place at a dose rate of approximately 1 Gy/min (18).

RNA Isolation

Total RNA for real-time quantitative PCR detection analysis was extracted from all cell lines using TRIzol reagent (Invitrogen) according to the instructions of the manufacturer. RNA concentrations were determined by absorbance at 260 nm, and RNA quality was verified by the integrity of 18 S and 28 S rRNA using ethidium bromide staining of total RNA samples subjected to 1.2% agarose gel electrophoresis. Synthesis of cDNA from total RNA (25 ng for microRNA and 100 ng for mRNA) was performed using an Applied Biosystems kit.

qRT-PCR cDNA Synthesis

Reverse transcription reactions were performed using the TaqMan microRNA reverse transcription kit (catalog no. 4366596, Invitrogen) with 100 ng of total RNA per 20-μl reaction. Reverse transcription thermocycling parameters were as follows: 16 °C for 30 min, 42 °C for 30 min, 85 °C for 5 min for miRNA and 25 °C for 10 min, 37 °C for 60 min, 37 °C for 60 min, and 85 °C for 5 min for mRNA. Reactions were performed on a MyCycler (Bio-Rad). Real-time PCR was performed using the ABI 7900HT system in 10-μl reaction volumes containing 1 μl of each primer, 5 μl of 2× iQ^TM SYBR Green Supermix, 2 μl of nuclease-free water, and 1 μl of cDNA (total 10 μl) in optical 384-well plates (Applied Biosystems). Cycling conditions were as follows: 95 °C for 10 min, followed by 40 cycles of 95 °C for 15 s and 60 °C for 60 s. Triplicate quantitative PCR reactions were performed for each cDNA sample for all experiments. The threshold fluorescence level was set manually for each plate using SDS software version 2.3 (Applied Biosystems). Following the export of cycle threshold (Ct) data, further data analysis for both platforms was performed in Microsoft® Excel 2003. Comparison of slope and R² values between preamplified and non-amplified cDNA, as a template on the BioMark arrays, was performed as a paired Student's t test in Microsoft® Excel 2003.

Cell Cycle Analysis

All cultures were incubated at 37 °C and 5% CO₂. Subsequently, parallel cultures were either exposed to 4 Gy IR (Rad-Source) or left untreated. Cells were returned to a 37 °C, 5% CO₂ incubator for 24 h and assayed for viability and DNA content. Cell viability was determined by trypan blue exclusion. For cell cycle analysis, cells were collected by centrifugation and resuspended at 1 × 10⁶ cells/ml in propidium iodide (PI) staining buffer (0.1% sodium citrate, 0.1% Triton X-100, and 50 mg/ml PI) and were treated with 1 mg/ml RNase at room temperature for 30 min. Cell cycle histograms were generated after analysis of PI-stained cells by FACS with a BD Biosciences FACScan. For each culture, at least 1 × 10⁴ events were recorded. Histograms generated by FACS were analyzed by ModFit cell cycle analysis software (Verity, Topsham, ME) to determine the percentage of cells in each phase (G₁, S, and G₂/M).

Colonogenic Survival Assay

Cell survival was evaluated by colony formation assay in HBECs (EV and 15b/16-2 stable lines) following IR at 4 Gy as described previously (19). Briefly, HBECs were transfected with pre-miR-15b and silencer-miR-15b and incubated for 24 h. Cells were further irradiated with 4 Gy and seeded for colony formation assays. The plates were left undisturbed, and, 3 weeks following treatment, colonies were fixed with 70% ethanol, stained with 1% methylene blue, and the number of positive colonies were counted (>50 cells). The survival fraction was calculated as follows: (number of colonies for treated cells / number of cells plated) / (number of colonies for corresponding control / number of cells plated). Experiments were performed in triplicate (20).

Immunofluorescence Staining

To assay DNA repair, 4 × 10⁴ cells were seeded on sterile coverslips. After 12 h, cells were treated with IR at 4 Gy. Cells were fixed with cold methanol at the indicated time points, and immunofluorescence was performed to detect Rad51 foci (20). To assay for DNA double strand breaks, 4 × 10⁴ cells were seeded on sterile coverslips and exposed to 4 Gy IR. Cells were fixed with cold methanol at the indicated time points, and γ-H2AX immunofluorescence was performed as described previously (21). Briefly, cells were rinsed in PBS and incubated for 5 min at 4 °C in ice-cold cytoskeleton buffer (10 mm Hepes/KOH (pH 7.4), 300 mm sucrose, 100 mm NaCl, and 3 mm MgCl₂) supplemented with 1 mm PMSF, 0.5 mm sodium vanadate, and proteasome inhibitor (Sigma, 1:100 dilution), followed by fixation in 70% ethanol for 15 min. The cells were blocked and incubated with anti-Rad51 (1:1000, Santa Cruz Biotechnology) or anti-γ-H2AX (1:1000, Millipore). The secondary antibody was anti-rabbit Alexa Fluor 594-conjugated or anti-mouse Alexa Fluor 488-conjugated antibody (1:2000, Invitrogen). DAPI (Invitrogen, catalog no. D21490) was used for nuclear staining. The coverslips were subsequently mounted onto slides with mounting medium (Aqua Poly Mount, Polysciences, Inc., catalog no. 18606) and analyzed via fluorescence microscopy (Carl Zeiss, Thornwood, NY). Positive and negative controls were included in all experiments. A total of 500 cells was assessed according to the standard procedure.

Western Blot Analysis

Western blotting was performed on the total protein extracts of the cell lines with and without 24 h of exposure to IR. The harvested cells were washed three times with ice-cold PBS and lysed in radioimmune precipitation assay buffer (150 mm NaCl, 50 mm Tris (pH 8.0), 5 mm EDTA, 0.5% sodium deoxycholate, 0.1% SDS, and 1.0% Nonidet P-40) with protease and phosphatase inhibitor mixtures (Sigma-Aldrich) for the total protein fraction. Protein concentrations in cell extracts were determined using the Bradford assay (Bio-Rad). 30 μg of total lysates were diluted 1:1 in radioimmune precipitation assay SDS-PAGE sample buffer, loaded onto 12% polyacrylamide gels, and blotted onto polyvinylidene difluoride membranes (Bio-Rad). Membranes were blocked with 5% nonfat milk in PBS (pH 7.6), 0.2% Tween 20 for 1 h and then incubated with anti-ATM (1:1000, Santa Cruz Biotechnology) mouse monoclonal antibody, anti-phospho-ATM (1:1000, Cell Signaling Technology) polyclonal rabbit antibody, anti-Chk1 (1:1000, Santa Cruz Biotechnology) mouse monoclonal antibody, anti-phospho Chk1 (1:1000, Cell Signaling Technology) rabbit polyclonal antibody, anti-phospho-p53 (1:1000, Cell Signaling Technology) mouse monoclonal antibody, anti-MDM2 (1:1000, Santa Cruz Biotechnology) rabbit polyclonal antibody, anti-p53 (1:1000, Santa Cruz Biotechnology) mouse polyclonal antibody, anti-BID (1:1000, Santa Cruz Biotechnology) rabbit polyclonal antibody, anti-Cyclin D1 (1:1000, Santa Cruz Biotechnology) mouse monoclonal antibody, anti γ-H2AX (1:1000, Millipore) mouse monoclonal antibody, anti-Rad51 (1:1000, Santa Cruz Biotechnology) rabbit polyclonal antibody, and anti-Wip1 (1:1000, Santa Cruz Biotechnology) rabbit polyclonal antibody. After washing in PBS (pH 7.6) and 0.2% Tween 20, the membranes were incubated with a horseradish peroxidase-conjugated goat anti-rabbit antibody (1:2000, Cell Signaling Technology) or anti-mouse antibody (1:2000, Cell Signaling Technology), and the immunoblots were visualized using ECL detection kits (Pierce). A mouse anti-β-actin antibody (1:2000, Santa Cruz Biotechnology) was used as the control for equal loading of total lysate.

RT Profiler PCR Array

RNA from EV and miR-15b/16-2 stable cells with and without IR was converted into cDNA using the RT² First Strand kit (SABiosciences). This cDNA was then added to the RT² SYBR Green quantitative PCR Master Mix (SABiosciences). Next, each sample was aliquotted on the DNA damage pathway PCR arrays (SABiosciences, catalog no. PAHS-029Z). All steps were done according to the protocol of the manufacturer for the ABI Prism 7000 HT detection system. To analyze the PCR array data, a Microsoft Excel sheet with macros was downloaded from the web site of the manufacturer. Expression fold differences were calculated using the −ΔΔCt method. β-Actin was used as an endogenous control to calculate ΔCt values (Ct_gene − Ct_β-actin). Gene expression fold differences were calculated relative to the averaged ΔCt value in 15b/16-2 and EV cell lines as the ratio of 2-ΔCt between the cell lines (2^{−ΔCt 15b/16-2 cell}/2^{−ΔCt EV cell}).

Target Analysis

Bioinformatics analysis was performed by using the following specific software programs: Targetscan, RNA22, and RNAhybrid. These programs identified the 3′UTRs of Wip1 as miR-15b binding sites.

Plasmid Construction

To generate Wip1 3′UTR luciferase reporter constructs, the 3′UTRs were amplified by PCR and cloned into psiCHECK2 vector at XhoI and NotI restriction sites (Promega). A mutant Wip1 3′UTR plasmid was also constructed in the psiCHECK2 vector by deleting the miR-15b targeting sequence in the 3′UTR region of Wip1.

Dual-Luciferase Reporter Assay

To confirm that miR-15b can bind to the 3′UTRs of Wip1, 2 × 10⁵ HEK293A and HBECs were seeded in 12-well plates with miR-15b precursors or scrambled sequence miRNA control (Ambion) overnight. Cells were then cotransfected with either WT Wip1 3′UTR psiCHECK2 or mutant (mt) Wip1 3′UTR psiCHECK2 reporters. Forty-eight hours after cotransfection, luciferase assays were performed using the Dual-Luciferase reporter assay kit (Promega). Firefly luciferase activity was normalized to Renilla luciferase activity for each sample. Statistical significance was analyzed by unpaired Student's t test.

Statistical Analysis

The focus staining calculation was done by Student's t test and survival fraction, and data were analyzed via analysis of variance followed by a Bonferroni post test using GraphPad Prism version 4.02 (GraphPad Software, San Diego, CA). Data are presented as mean ± S.E. SA Biosciences PCR arrays were calculated using web-based analysis software.

RESULTS

Regulation of miR-15b in HBECs Is Mediated by Ionizing Radiation

Patterns of miR-15b expression levels were measured in HBECs in response to low and high doses of IR (Rad-Source). miR-15b expression levels were induced by IR (Fig. 1A) in a time- and dose-dependent manner (Fig. 1A). Two hours after IR, miR-15b expression reached its peak in all doses compared with other time points. Following the peak at 2 h, the miR-15b expression level decreased gradually and, by 24 h, had returned to baseline levels (Fig. 1A).

FIGURE 1.

IR induces miR-15b expression in a time- and dose-dependent manner. A, effect of x-ray-irradiation on the expression levels of miR-15b in HBECs. In all different doses examined, miR-15b expression was highest at the 2 h time point and decreased gradually over 24 h. B, HBECs were transfected with pre-miR-15b and silencer-miR-15b and incubated for 24 h. Cells were further irradiated and seeded for colony formation assays. miR-15b promoted colony formation of HBECs. Data are the mean survival fraction ± S.E. from at least three independent colony formation assays following IR. *, p < 0.001. C, HBECs, EV, and miR-15b/16-2 stable cells were irradiated (4 Gy) and seeded for a colony formation assay. 15b/16-2 stable cells displayed more colony formation compared with HBECs and EV cells. Data are the mean survival fraction ± S.E. from at least three independent experiments. *, p < 0.001.

miR-15b Promotes Survival in HBECs Exposed to IR

Following exposure to IR (Rad-Source), we observed that miR-15b overexpression significantly promoted HBEC survival. As seen in Fig. 1B, the combination of pre-miR-15b transfection and IR led to an increase in the colony-forming ability of HBECs compared with knockdown miR-15b transfection with IR. We also observed an increase in survival after IR treatment in 15b/16-2 stable cells compared with EV and non-transfected HBECs (Fig. 1C).

miR-15b/16-2 Regulates IR-induced DNA Damage and Repair

We observed that miR-15b expression is normally increased in HBECs following IR. However, the biological relevance of this alteration remains unclear. Both EV and 15b/16-2 stable cells were exposed to IR and assessed for DNA damage. We measured the number of γ-H2AX foci (an in situ marker of double strand breaks) 2 h after exposure to IR. As shown in Fig. 2A, there was a significant increase in γ-H2AX foci in miR-15b/16-2 stable cells compared with EV cells, indicating a substantial increase in double strand breaks. We also examined for DNA repair through the measurement of Rad51 foci, a well established functional marker of DNA repair activity, 5 and 12 h following IR (Fig. 2C). Rad51 focus expression was significantly higher in 15b/16-2 stable cells at 5 h compared with EV cells, whereas the expression for both cell lines was very low by 12 h following IR. With 4-Gy treatment, there was only a modest increase in survival with miR-15b/16-2 overexpression (Fig. 2C). DNA damage (Fig. 2B) 2 h after IR and repair (Fig. 2D) 5 h after IR were also evaluated by Western blot analysis. These results suggest that miR-15b and 16-2 contribute to both DNA damage and repair.

FIGURE 2.

miR-15b/16-2 regulates IR-induced DNA damage and repair. EV and 15b/16-2 stable cells were treated with 4-Gy IR. Cells were then processed for immunofluorescence staining for γ-H2AX foci after 2 h of IR and Rad51 foci after 5 and 12 h of IR. A total of 500 cells were counted in each group under fluorescence microscopy. More than 10 foci/cell were considered to be positively stained cells. Data are representative of three independent experiments the % of cells (mean ± S.E.; *, p < 0.001) >γ-H2AX foci (A), with >10 Rad51 foci (C). Also shown is representative staining of cells exhibiting γ-H2AX foci (A) foci and Rad51 foci (C) (green), with the nuclei stained with DAPI (blue). Cells were also processed for WB with γ-H2AX (B) and Rad51 (D).

15b/16-2 Cells Undergo Cell Cycle Arrest and Apoptosis after IR

We next evaluated whether IR had any effect on the cell cycle of miR-15b/16-2 stable cells. When irradiated with 4 Gy, miR-15b/16-2 cells arrested largely in G₂/M1 phase (53.1%), with much less in S (7.2%) and G₁/G₀ (34.1%) with respect to EV cells (Fig. 3, A and B). We next sought to determine whether these observed effects on the cell cycle were mediated by miR-15b. We did not observe any difference in the distribution of the cell cycle in IR-treated HBECs following miR-15b knockdown (Fig. 3, C and D). These results suggest that miR-15b knockdown was insufficient to reverse the observed effects of miR-15b gain of function. Next, we observed that irradiated miR-15b/16-2 cells underwent apoptosis on the basis of PARP cleavage 48 h following IR (Fig. 3E). We next evaluated the expression of common regulators of G₂/M1 cell cycle progression. Synchronized miR-15b/16-2 stable and EV cells were treated with 4-Gy irradiation, and lysates were subjected to Western blotting with specific antibodies.

FIGURE 3.

IR-treated 15b/16-2 cells show increased arrest in G₂/M phase as well as induced apoptosis. miR-15b/16-2 and EV stable cells were irradiated (4 Gy) and cultured for an additional 24 h. Cells were harvested and then processed to determine the DNA content by PI staining and flow cytometry. A, representative images of the cell cycle distribution of EV and 15b/16-2 cells with and without IR. B, graphical representation of the percentage of cells in different phases of the cell cycle. HBECs were transfected with silencer-miR-15b or scramble miR and incubated for 24 h. Cells were irradiated (4 Gy) and cultured for another 24 h. Cells were harvested and then processed to determine the DNA content by PI. C, representative images of the cell cycle distribution of HBEC-miR-15b K_D and HBEC-scramble cells with and without IR. D, graphical representation of the percentage of cells in the different phase of cell cycle. 15b/16-2 cells induce cell cycle arrest in G₂/M₁ phase after IR. Following 48 h of IR treatment, cell lysate was prepared, and the Western blot was assessed for apoptosis (E) using a cleaved PARP antibody.

IR Induced Phosphorylation of ATM/Chk1/p53 and Inhibited Cyclin D1 Expression in 15b/16-2 Cells

Phosphorylated ATM, Chk1, phosphorylated p53, and BID expression were up-regulated in 15b/16-2 cells compared with EV (Fig. 4A) when treated with IR. We did not observe any baseline expression of phosphorylated ATM, Chk1, and p53 in the setting of no IR (Fig. 4B) in both EV and 15b/16-2 stable cells. MDM2 expression was down-regulated in 15b/16-2 stable cells compared with EV cells when treated with IR (Fig. 4A). Cyclin D1 expression was down-regulated in 15b/16-2 cells treated with IR, as shown in Fig. 4A. Without IR, there was no change of Cyclin D1 expression in both cell lines (Fig. 4B). We examined whether BID was modified in response to DNA damage and found that BID expression was increased in miR-15b/16-2 cells (Fig. 4, A and B). We also conducted a time course for phospho-p53 and MDM2 protein expression in 15b/16-2 and EV cells. Phospho-p53 expression was higher in 15b/16-2 cells from 2–48 h after IR compared with EV cells (Fig. 4C). On the other hand, MDM2 protein expression was higher in EV cells from 2–48 h compared with 15b/16-2 cells (Fig. 4C). To further validate the relationship between miR-15b, p-p53, and MDM2, we knocked down miR-15b and examined the p-p53 and MDM2 expression in HBECs. We observed that miR-15b knockdown increased MDM2 expression (Fig. 4D). These results suggest that miR-15b may contribute to the induction of phosphorylated ATM and p53 expression and down-regulated MDM2 expression in the setting of IR exposure. Conversely, knockdown of miR-15b in HBECs resulted in down-regulation of γ-H2AX and Rad51 expression (Fig. 4D). Knockdown of miR-15b in HBEC significantly reduced miR-15b expression following treatment with IR (Fig. 4E).

FIGURE 4.

IR induced phosphorylation of ATM/Chk1/p53 and inhibited Cyclin D1 expression in 15b/16-2 cells. Both EV and 15b/16-2 stable cells were treated with IR (4 Gy) and then cultured for 24 h. Total proteins were isolated using radioimmune precipitation assay lysis buffer, and Western blot analysis was performed using 20 μg of protein/lane. Shown is phosphorylated ATM, Chk1, MDM2, p53, total p53, BID, and Cyclin D1 expression following treatment with IR (A) and without IR (B). C, time course for phosphorylated p53 and MDM2 expression in EV and 15b/16-2 cells at 0, 2, 4, 24, and 48 h with 4-Gy IR. D, HBECs were transfected with either scramble or silencer-miR-15b. 24 h after transfection, cells were treated with IR (4 Gy), and, 24 h later, total proteins were isolated and examined for total ATM, phosphorylated ATM, MDM2, total p53, phosphorylated p53, γ-H2AX, and Rad51 protein expression. E, HBECs were transfected with scramble and silencer-miR-15b and incubated for 24 h. Cells were then treated with IR (4 Gy) and incubated for another 24 h. Total RNA was extracted, cDNA was synthesized, and qRT-PCR was performed with miR-15b primer (*, p < 0.001).

Global Profiling Demonstrates That miR-15b/16-2 Induces ATM/ATR, Cell Cycle, and DNA Repair Pathway Genes following Irradiation

We conducted a global analysis for genes participating in DNA damage signaling, cell cycle arrest, and DNA repair in both EV and miR-15b/16-2 stable cells with and without IR using a quantitative PCR array (supplemental Tables 1 and 2). We grouped genes according to their pathways: ATM/ATR signaling pathway, apoptosis, cell cycle, and DNA repair. In ATM/ATR signaling, nine genes were analyzed, and one-third of genes were up-regulated (PARP1, TP53, and MDC1), whereas 22% of genes were down-regulated (RAD1 and RNF8) in miR-15b/16-2-overexpressing cells compared with EV without IR. These results suggest that miR-15b, in combination with IR, induced an increase in the expression levels of ATM/ATR and DNA repair signaling genes (supplemental Tables 1 and 2).

Wip1 (PPM1D) Is a Target of miR-15b/16-2

We searched for potential targets of miR-15b using three in silico target prediction programs: Targetscan, RNA22, and RNAhybrid. We identified Wip1 (PPM1D) as a target for miR-15b (Fig. 5A). Using the 3′UTR both with and without mutations, we validated Wip1 as a target for miR-15b (Fig. 5B). Next, we examined the relationship between miR-15b and Wip1 expression at the mRNA level. miR-15b overexpression led to a reduction in Wip1 mRNA, whereas miR-15b knockdown resulted in an increase in Wip1 mRNA (Fig. 5, C and D). In the setting of IR exposure, we observed an early induction of miR-15b at 2 h, whereas, 24 h after IR, miR-15b expression levels were reduced significantly (Fig. 5E). On the other hand, Wip1 expression levels increased at 24 h (Fig. 5F). We also observed that miR-15b overexpression reduced Wip1 at the protein level (Fig. 5G), suggesting that miR-15b and Wip1 are inversely correlated.

FIGURE 5.

Wip1 is a target of miR-15b. A, predicted binding sites of hsa-miR-15b to the 3′UTR of Wip1 (PPM1D) as detected by three separate in silico prediction programs. Wip1 3′UTR (WT) contains a predicted miR-15b binding site that is aligned with miR-15b sequence. A mutant Wip1 3′UTR (mt) plasmid was constructed by deleting the underlined sequence. B, luciferase reporter constructs containing Wip1 3′UTR (WT)/Wip1 3′UTR (mt) were cotransfected with scrambled or pre-miR-15b into HBEC. Relative firefly luciferase expression was normalized to Renilla luciferase. Data are representative of three independent experiments (mean ± S.E.; *, p < 0.001). C and D, HBECs were transfected with either precursor-miR-15b or knock-down miR-15b. 24 h after transfection, RNA was extracted, cDNA was synthesized, and RT-PCR was conducted to detect Wip1. HBECs were treated with IR (4 Gy), and, at the indicated time points, RNA was extracted, cDNA was synthesized, and RT-PCR was performed for miR-15b (E) and Wip1 (F). G, both EV and 15b/16-2 stable cells were irradiated, and, after 24 h, total proteins were isolated and examined for the presence of Wip1.

Wip1 Re-expression Results in a Partial Reversal of miR-15b/16-2-mediated Effects on DNA Damage and Response

To determine whether the observed effects of miR-15b/16-2 on IR DNA damage and response were mediated by targeting of Wip1, we rescued Wip1 expression in miR-15b/16-2 stable cells by transfecting them with FLAG tag Wip1 lacking the 3′UTR. 24 h after transfection, cells were exposed to IR and incubated for another 24 h. Total proteins were extracted, and the presence of Wip1, total and phosphorylated ATM, total and phosphorylated Chek1, MDM2, phospho-p53, γ-H2AX, Rad51, and cleaved PARP was assessed by Western blot analysis (Fig. 6A). We first confirmed the overexpression of Wip1. We observed that Wip1 rescue in miR-15b/16-2 stable cells led to an increase in MDM2 protein levels and decreased expression of phosphorylated p53, γ-H2AX, Rad51, and cleaved PARP (Fig. 6A). We did not observe significant baseline expression of phosphorylated ATM, Chk1, γ-H2AX, Rad51 and cleaved PARP in the setting of no IR treatment (Fig. 6B). These results suggest that the effects of miR-15b are partially mediated through targeting of Wip1. We propose a scheme for miR-15b involvement as a mediator of IR response in HBECs (Fig. 6C).

FIGURE 6.

Wip1 re-expression results in a partial reversal in miR-15b/16-2-mediated effects on DNA damage and response. 15b/16-2 stable cells were transfected with a Wip1FLAG plasmid lacking a 3′UTR. 24 h after transfection, cells were treated with IR (4 Gy) or left untreated and incubated for another 24 h, after which total proteins were extracted. The presence of Wip1, total ATM, phosphorylated ATM, total Chk1, phosphorylated Chk1, MDM2, phosphorylated p53, g-H2AX, Rad51, and C-PARP was verified by Western blot treated with IR (A) or without IR (B). C, we hypothesize that IR-induced DNA damage increases miR-15b expression, which, in turn, induces phosphorylation of p53 directly or via targeting of Wip1. Wip1 inhibits phosphorylation of p53 directly or via inhibition of MDM2, leading to an induction in apoptosis, cell cycle arrest, and DNA repair.

DISCUSSION

Acute radiation pneumonitis and chronic radiation fibrosis represent common consequences of radiation exposure in the setting of treatment of various cancers (22), including non-small cell lung cancer, small cell lung cancer (23), breast cancer (24), esophageal cancer (25), mediastinal tumors (26), and head and neck tumors (27). Although the underlying biology of radiation-induced injury to the lung is becoming increasingly known, few actionable targets exist for the purposes of therapy. In this study, we examined the role miRNAs as potential mediators of radiation-induced injury to the airway epithelium.

miRNAs and their processing have emerged rapidly, critically contributing to both lung development and the pathogenesis of various diseases of the lung. For example, mice with conditional knockout of the key processing enzyme Dicer in lung epithelial cells exhibit a failure of epithelial branching, underscoring the essential regulatory role of miRNAs in lung epithelial morphogenesis (28). Several studies are ongoing to identify clinically relevant biomarkers in lung disease. In this study, we attempted to identify the mechanisms by which miR-15b may serve as a mediator of the DNA damage response. This is largely on the basis of previous observations that miR-15b is deregulated following exposure to various genotoxic stressors as well as in the setting of sepsis. Recently, it has been reported that members of the miR105/107 family (miR-15b, miR-424, and miR-107) were all up-regulated, with miR-15b more abundantly localized in areas of emphysema as well as fibrosis (29). However, the significance of this observation is unknown. We hypothesized that such findings may suggest that miR-15b could serve as a marker of injury or important to maintaining homeostasis between injury and repair. Our findings suggest that miR-15b may play an important role in DNA damage response and repair mechanisms and, therefore, protect cells from genotoxic stress.

Many mechanisms are induced upon DNA damage, including altered gene expression, activation of cell cycle checkpoints, and elevated DNA repair activity, all with a goal of maintaining genomic integrity. This is an area of intense investigation. Our results suggest that miR-15b deregulation could be an important factor contributing to the DNA damage-induced cellular response. We detected increased expression of H2AFX, RAD1, TP53, CDC25A, MDC1, EXO1, UNG, BRCA1, and PARP1 genes in miR-15b/16-2 overexpressing HBECs (supplemental Tables 1 and 2). These genes have been implicated as regulators of the ATM/ATR signaling, apoptosis, cell cycle, and DNA repair pathways. Although the majority of our studies were conducted using a miR-15b/16-2-overexpressing cell line, we also observed that selected gain of function or knockdown of miR-15b could alter the cell phenotype and selected targets. We demonstrated that miR-15b/16-2 is a key switch in ATM/Chek1/p53 signaling in accelerating the DNA damage response and repair in the setting of IR exposure. Both ATM and Chk1 have been reported to directly phosphorylate p53 on several N-terminal sites (30). It has also been reported that IR activates p53 by inducing phosphorylation at the N- and C-terminal phosphorylation sites as well as direct down-regulation of MDM2 expression (31). We observed a time-dependent induction in phosphorylated p53 and decrease in MDM2 levels following IR in miR-15b/16-2 cells. Conversely, miR-15b knockdown led to an increase in MDM2 protein and reduction in p53 phosphorylation. These results suggest a direct role of miR-15b in the relationship between p53 and MDM2. ATM kinase plays a pivotal role in the immediate response of cells to double strand breaks. It has been reported that ATM kinase is important in the process by which IR induces phosphorylation of BID, which plays an important role in cell cycle arrest in G₂/M phase rather than apoptosis (32). miR-15b-induced G₂/M cell cycle arrest may facilitate the DNA repair process, contributing to genomic stability in stressed cells. It has been reported that γ-H2AX foci were increased in IR-induced G₂/M cell cycle arrest (33). We also observed γ-H2AX expression in HBECs with miR-15b knockdown treated with IR (Fig. 4D). Our results suggest that miR-15b contributes to the induction of γ-H2AX expression following IR exposure. To our knowledge, this represents the first report that miR-15b/16-2 can induce γ-H2AX, a known DNA damage marker, and Rad51, a known marker for DNA repair in human bronchial epithelial cells.

Wip1 is a serine/threonine phosphatase that has been shown to inactivate the DDR through several mechanisms, including dephosphorylation of p53, γ-H2AX, and ATM. Here we identified Wip1 as a direct and functional target of miR-15b. Furthermore, we observed that reintroduction of Wip1 in miR-15b/16-2-overexpressing HBECs led to a partial reversal in the DDR in the setting of IR exposure.

In conclusion, our study suggests that DNA damage by irradiation induces miR-15b expression, causing up-regulation of ATM/Chk1. miR-15b/16-2 may phosphorylate p53 via ATM/Chek1 or by inhibiting Wip1. We hypothesized that miR-15b inhibition of Wip1 leads to a down-regulation of MDM2 and a resultant increased phosphorylation of p53 (Fig. 6B). Therefore, p53 stabilization may inhibit Cyclin D1 and induce apoptosis or DNA repair as well as cell cycle arrest. This study further suggests that the link between miR-15b/16-2 and DNA repair may serve as a mechanism for protection of the lung from cellular stress. Further studies are required to establish the mechanisms of regulation of miR-15b and 16-2 in the setting of cellular stress, with the eventual goal of developing new clinical biomarkers and actionable targets in radiation-induced injury to the lung.