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. Author manuscript; available in PMC: 2022 Sep 2.
Published in final edited form as: DNA Repair (Amst). 2019 Mar 23;78:37–44. doi: 10.1016/j.dnarep.2019.03.012

Ionizing radiation-induced growth in soft agar is associated with miR-21 upregulation in wild-type and DNA double strand break repair deficient cells

Siyuan Tang a,b, Bailong Liu a,c, Min Liu a,c, Zhentian Li a, Jiaqi Liu a, Hongyan Wang a, Jian Wang a, You-Take Oh a, Liangfang Shen b,*, Ya Wang a,*
PMCID: PMC9437867  NIHMSID: NIHMS1626762  PMID: 30954901

Abstract

DNA double strand breaks (DSBs) are a severe threat to genome integrity and a potential cause of tumorigenesis, which is a multi-stage process and involves many factors including the mutation of oncogenes and tumor suppressors, some of which are transcribed microRNAs (miRNAs). Among more than 2000 known miRNAs, miR-21 is a unique onco-miRNA that is highly expressed in almost all types of human tumors and is associated with tumorigenesis through its multiple targets. However, it remains unclear whether there is any functional link between DSBs and miR-21 expression and, if so, does the link contribute to DSB-induced genomic instability/tumorigenesis. To address this question, we used DNA-PKcs−/− (deficient in non-homologous end-joining (NHEJ)) and Rad54−/− (deficient in homologous recombination repair (HRR)) mouse embryonic fibroblasts (MEFs) since NHEJ and HRR are the major pathways for DSB repair in mammalian cells. Our results indicate that levels of miR-21 are elevated in these DSB repair (DSBR) deficient cells, and ionizing radiation (IR) further increases these levels in both wild-type (WT) and DSBR-deficient cells. Interestingly, IR stimulated growth in soft agar and this effect was greatly reduced by blocking miR-21 expression in both WT and DSBR-deficient cells. Taken together, our results suggest that either IR or DSBR-deficient can lead to an upregulation of miR-21 levels and that miR-21 is associated with IR-induced cell growth in soft agar. These results may help our understanding of DSB-induced tumorigenesis and provide information that could facilitate the development of new strategies to prevent DSB-induced carcinogenesis.

Keywords: DNA double strand breaks, miR-21, non-homologous end-joining, homologous recombination repair, soft agar, ionizing radiation

1. Introduction

Among more than 2000 human microRNAs (miRNAs), miR-21 is unique since it overexpresses in almost all types of human tumors/cancers (including brain, liver, colorectal tumors, as well as breast, pancreatic and lung cancers, etc.) [1, 2], and is associated with the promotion of tumorigenesis [3–5]. MiR-21 targets multiple tumor suppressors and inhibits their expression through binding to a partially matched sequence in their 3’ untranslated region; its targets include PTEN, PDCD4, SPRY2, TPM1, ANP32A, SMARCA4, p53, etc. [6]. We previously reported that ionizing radiation (IR) stimulates miR-21 expression by activating the AP-1 and EGFR pathways in human liver cells [3]. IR induces different types of damage in which SSBs and base damage predominate versus DSBs that are comparatively rare (in one mammalian cell, 1 Gy dose produces about 1000 SSBs, 2000 instances of oxidative base damage, and 20 DSBs [7]). Although it is known that single strand breaks (SSBs) and oxidative base damage are associated with stimulation of miR-21 expression [8–10], it remains unclear whether double strand breaks (DSBs) are also linked to stimulating miR-21 expression.

This study is to address the relationship between DSBs and miR-21 through three aims. The first aim is to determine whether DSBs are linked to miR-21 upregulation. We compared miR-21 levels in wild-type (WT) versus DSB repair (DSBR)-deficient mouse embryonic fibroblasts (MEFs). These DSBR-deficient MEFs lacked either DNA-PKcs that is required for non-homologous end joining (NHEJ) [11], or Rad54 that is required for homologous recombination repair (HRR) [12]. It is assumed that these DSBR-deficient cells have the normal ability to repair SSB and base damage since DSBR, SSBR and base damage repair use different pathways involving different proteins [13–15], and no reports show abnormal repair of SSBs or base damage in these DSBR-deficient cells. The second aim is to determine whether inhibiting miR-21 expression reduces DSB-promoted cell growth in soft agar since cell growth in soft agar to form colonies is commonly used in vitro to detect the ability of anchorage-independent growth (oncogenic transformation), a hallmark of carcinogenesis [16]. The third aim is to explore whether IR-activated ATM and ATR, are also associated with miR-21 upregulation.

Our results suggest that IR, including endogenous DSBs, are associated with EGFR-dependent miR-21 upregulation. Additionally, inhibition of miR-21 reduces soft agar colony-forming efficiency of irradiated cells, which is more clearly observed in DSBR-deficient cells. Finally, we show that inhibition of IR-activated ATM and ATR also mildly reduce miR-21 expression, which might be related to their effects on EGFR activation and AP-1 expression. Taken together, these results may not only help our understanding of the mechanism underlying DNA DSB-induced tumorigenesis, but may also facilitate the development of new strategies to prevent tumorigenesis.

2. Materials and methods

a. Cell lines and irradiation

All of the MEFs used in this study were derived from C57BL/6J mice. Rad54−/− (HRR deficient) and their WT control MEFs were obtained from Dr. George Iliakis’s lab after obtaining approval from Dr. Kevin Mills [17]. DNA-PKcs−/− (NHEJ-deficient) MEFs were obtained from Dr. David Chen’s lab [11]. Additional WT MEF lines were obtained from the labs of Drs. Chuan-Yuan Li [18] and Gloria Li, respectively [19]. MiR-21 knock-in MEFs that continuously over-express miR-21 were derived from our lab (generated by GenOway Inc. France) as described previously [20, 21]. All the MEFs were grown in DMEM supplemented with 10% bovine calf serum. Radiation was performed using an X-ray machine (X-RAD 320, N. Branford, CT, USA) in our laboratory. The energy setting for the experiments was 320 kVp, 10 mA, with 2 mm aluminum filtration. The IR dose rate was 2 Gy/min, which was controlled by a computer with relevant software.

b. Quantitative real-time PCR measures miRNA or Egfr levels in cells

Total RNA was extracted from MEF cells using miRNeasy mini kits (#217004, Qiagen, Valencia, FL, USA). cDNA was synthesized using 500 ng RNA with the SuperScript VILO cDNA Synthesis Kit (#11754050, Life Technologies Scientific Corp, Carlsbad, CA, USA). The samples were performed in triplicate qPCR reactions using Fast SYBR™ Green Master Mix (#4385612, Thermofisher, Waltham, MA, USA) on an Applied Biosystems 7500 Fast real-time PCR system. Egfr mRNA level was measured using a forward primer, AGGCACAAGTAACA GGCTCAC, and a reverse primer, AAGGTCGTAATTCCTTTGCAC. Actg2 mRNA level (an internal control for analyzing Egfr mRNA level) was measured using a forward primer, TTGAACATGGCATTGTTACCAA and a reverse primer, TGGCATAGAGGTCTTTACGGA. A TaqMan miRNA reverse transcription kit (#4366596 Thermofisher, Waltham, MA, USA) was used to prepare the products for the assays (#4427975, Thermofisher, Waltham, MA, USA). All procedures were followed according to the manufactures’ instructions. The relative expression level of miRNA (miR-21, miR-155, miR-34a) was normalized to SNO202 using a 2ΔΔCT method [22].

c. siRNA treatment and other reagents

Pools of small interfering RNA (siRNA) oligonucleotides directed against Rad54 (sc-36363), DNA-PKcs (sc-35201), or STAT3 (sc-29493) and control RNA (sc-37007) were purchased from Santa Cruz Biotech Inc. (Santa Cruz, CA, USA). Human (Hsa)-miR-21–5p (same sequence as mouse miR-21–5p) inhibitor was purchased from Ambion Inc. (Austin, TX). WT MEFs (control for Rad54 deficient cells (shown in Fig. 2a) and an additional WT line obtained from Dr Gloria Li’s lab (shown in Supplementary Fig. 2)) were transfected with oligonucleotides using Lipofectamine 3000 (Invitrogen, San Diego, CA, USA) according to the manufacturer’s instructions. Cells were then collected at 24–48 h after transfection and whole cell lysates were prepared in RIPA buffer as described previously [3]. Protein levels were determined using antibodies (against Rad54 (D-18), DNA-PKcs (H-163), EGFR (sc-03), STAT3 (sc-482) or actin (sc-47778)) that were purchased from Santa Cruz Biotech Inc. Antibodies against AP-1 (c-Jun) (#9165), phosphorylated EGFR (#3777) and ATM (#2873) were purchased from Cell Signaling Technology Inc. (Danvers, MA, USA). A phosphorylated ATR antibody (ABE462) was purchased from Millipore-Sigma Inc. (Billerica, MA, USA). The ATM inhibitor (KU-55933 (S1092)), ATR inhibitor (VE821 (S8007)), and EGFR inhibitor (gefitinib (ZD1839)) were purchased from Selleckchem (Houston, TX, USA).

Fig. 2. High miR-21 and EGFR levels in Rad54−/− or DNA-PKcs−/− MEF cells are due to a deficiency in the DSBR gene.

Fig. 2.

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a. Effects of Rad54 or DNA-PKcs siRNA on the EGFR level in the WT control (for Rad54-deficient MEFs). At 48 h after transfection the cells were collected and whole cell lysates were prepared as described in Materials and methods. Actin was used as an internal loading control. b. Quantification of immunoblotting data (EGFR as described in (a)) and compared with miR-21 levels that were analyzed by qPCR. Data are normalized to control RNA-treated cells. Graph depicts the mean and standard deviation from three independent experiments; *, P<0.05; **, P<0.01. c. WT control (for Rad54-deficient MEFs), DNA-PKcs−/− or Rad54−/− cells were treated with a control or specific siRNA against Stat3 for 24 h, then exposed to IR (1 Gy) for an additional 24 h. Cells were then collected and cell lysates from a portion of the collected cells were prepared for detection of STAT3 levels using western blot. d. RNA was extracted from the remainder of the collected cells for detection of miR-21 levels. The miR-21 levels in cells treated with siRNA against Stat3 were normalized to the levels in WT cells treated with a control RNA (defined as “1.0”). The graph depicts the mean and standard deviation from three independent experiments; *, P<0.05. PK, DNA-PKcs−/− cells; R54, Rad54−/− cells.

d. Cell synchronization and flow cytometry

Cells were treated with nocodazole (Sigma-Aldrich, St. Louis, MO, USA) at a final concentration of 0.4 μg/mL for 20 h to the G2/M phase. Cells were subjected to a double-thymidine blockade for 14 h and released for 2 h to the S phase, as described previously [23]. The cell cycle distribution was examined using flow cytometry as described previously [23].

e. Foci assay

The γ-H2AX and 53BP1 foci assays [24–26], as well as XRCC1 foci assay [27] were performed as described, with minor modifications. Briefly, cells were grown on coverslips, then fixed in 4% paraformaldehyde for 15 min, permeabilized for 5 min on ice in 0.2% Triton X-100, and blocked in 10% normal goat serum. The coverslips were incubated with an γ-H2AX (#05–636) (Millipore), 53BP1 (ad175433) or XRCC1 (ab1838) (Abcam Inc. Cambridge, MA, USA) antibody for 1 h respectively, washed in PBS and 1% BSA, and incubated using an Alexa Fluor 488-conjugated goat anti-mouse or anti-rabbit secondary antibody (Invitrogen) for 1 h at room temperature. Cells were washed in PBS and mounted using Vectashield mounting medium with 4,6-diamidino-2-phenylindole (Vector Laboratories Inc. Burlingame, CA, USA). Fluorescence images were captured using an AxioScope A1 Epifluorescence microscope (Germany) equipped with an MRm Cooled Digital Camera and Axiovision software (version 4.8). Co-localization of γ-H2AX and 53BP1 foci positive cells were counted as DSB-containing cells.

f. IR-promoted cell growth in soft agar

As described in our previous publications [28–30], cell growth (to form colonies) in soft agar was measured. To increase IR-induced cell oncogenic transformation propensity, we continually sub-cultured MEFs for 2 months after exposure to 1 Gy IR. Our unpublished findings (confirmed through personal communication with Drs. Story and Ding at UT Southwestern Medical Center) have shown that maintaining the cells in culture for 2–3 months after IR exposure (2 months for mouse cells and 3 months for human cells), enhances the propensity of IR-induced cell oncogenic transformation. Low melting temperature agarose (4%) and 2 x culture medium were mixed to obtain a 0.75% agarose concentration. After 2 months of subculture, the cells were treated with 20 nM control RNA (non-specific oligonucleotides) or a miR-21 inhibitor (miR-21 antisense oligonucleotides) for 24 h, then cells were harvested and divided into three portions to measure (i) miR-21 levels, (ii) plating efficiency in touch-culture dishes and (iii) cell growth in soft agar. During (ii) and (iii) colony formation experiments, we added an additional control RNA or miR-21 inhibitor to medium or to soft agar until colony formation ended. MiR-21 levels were measured using qPCR as described above. For anchorage-dependent growth, cells were harvested and 200 cells in 4 mL medium containing 20 nM miR-21 inhibitor or control RNA were plated. The cells were cultured at 37°C with 5% CO2 for 1 week. Colonies were stained with 0.2% p-iodonitrotetrazolium violet and >30 cell-colony was counted. For anchorage-independent growth, 5000 cells (2000 cells for miR-21 knock-in MEF) in 2 mL medium (containing freshly added miR-21 inhibitor or control RNA) were mixed with 2 mL 0.75% agarose (final 0.35% agarose and 20 nM miR-21 inhibitor or control RNA). The cells were cultured at 37°C with 5% CO2 for 3 weeks. The soft agar cultures were solidified at 4°C, stained with 0.2% p-iodonitrotetrazolium violet (Millipore-Sigma), and scanned for colony counting (colonies larger than 100 μm in diameter were included). The data were normalized using the plating efficiency (anchorage-dependent growth colonies).

g. Statistical analysis

Differences in protein/miRNA levels and soft agar colony forming efficiencies between two groups (comparing the values from one cell line with or without treatment; or comparing the values from two cell lines (WT and Rad54−/− or WT and DNA-PKcs−/−) with the same treatment) were evaluated using Pearson chi-square test or Student’s t-test. P values < 0.05 were regarded as significant.

3. Results

a. IR enhanced EGFR-dependent miR-21 upregulation in DSBR-deficient cells

To study whether DSBs alone are associated with miR-21 upregulation, we chose DNA-PKcs−/− (NHEJ-deficient) and Rad54−/− (HRR-deficient) MEFs since NHEJ and HRR are the two major DSBR pathways in mammalian cells. Notably, even without IR, DNA-PKcs−/− or Rad54−/− cells showed higher levels of miR-21 compared to WT cells (Fig. 1a, Supplementary Fig. 1a), suggesting that endogenous DNA DSBs may be associated with stimulation of miR-21 expression. IR enhanced the phenotype (Fig. 1a). EGFR, the miR-21 upstream regulator, increased its level in DNA-PKcs−/− or Rad54−/− (DSBR-deficient) cells versus in WT cells even without IR (Fig. 1b, c, Supplementary Fig. 1b), strongly supporting this. To study whether the relatively higher EGFR level in the DSBR-deficient cells was a common phenotype, we compared EGFR levels of the DSBR-deficient cells and three different WT MEF lines that were obtained from three different labs. All WT MEF lines showed a relatively lower EGFR level than the DSBR-deficient cells (Supplementary Fig. 1c), confirming that the DSBR-deficient cells do express a higher EGFR level compared to WT MEFs. To further verify the results, we quantified EGFR levels of WT and DSBR-deficient cells, which were shown on the same gel. The results clearly indicated higher EGFR levels in DSBR-deficient cells and IR enhanced the phenotypes (Fig. 1c), providing additional evidence to support the association of DSBs with EGFR-dependent miR-21 upregulation.

Fig. 1. Rad54−/− or DNA-PKcs−/− MEFs showed higher levels of spontaneous and IR-promoted upregulation of miR-21 and EGFR.

Fig. 1.

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a. MiR-21 levels determined by qPCR in MEFs (WT, Rad54−/− or DNA-PKcs−/−) at different times after exposure to 1 Gy IR. Graph depicts the mean and standard deviation of nine samples from three independent experiments. *, P<0.05. **, P<0.01; the data represent significant difference at each time point between WT and Rad54−/− cells or WT and DNA-PKcs−/− cells. b. EGFR levels in different MEFs measured at different times after 1 Gy IR by western blot (WT control for Rad54-deficient cells was used in the experiments). Actin was used as an internal loading control. Similar results were obtained from three independent experiments including a WT control for Ku80 deficient cells). c. Left panel: EGFR levels in different MEFs measured at 6 h after 1 Gy of IR by western blot shown in the same gel. Right panel: Quantification of the immunoblotting data shown in left panel. *, P<0.05. **, P<0.01. d. Left panel: image of γ-H2AX foci positive cells (scale bar = 5 μm); right panel: percentage of γ-H2AX foci positive cells in different MEFs at different times after IR (1 Gy). γ-H2AX foci positive cells were analyzed from 100 cells at each time point. Data represent mean ± s.d. from three independent experiments, *, P<0.05. **, P<0.01 (data were compared between WT and Rad54−/− or WT and DNA-PKcs−/− cells).

To examine whether the increased miR-21 levels in DSBR-deficient cells vs WT cells were also involved in SSBs, we examined XRCC1 foci in irradiated WT and DSBR-deficient cells using the approach as described [27] since such foci represent activated SSB repair (SSBR) status [31]. Although WT and DSBR-deficient cells showed clear IR-induced XRCC1 foci, these cells did not show any clear difference in XRCC1 foci at different times after IR (data not shown), supporting that SSBR is functional in the DSBR-deficient cells, and SSBs are not involved in the increased miR-21 levels of DSBR-deficient cells compared to that of WT cells. In contrast, γ-H2AX foci (DSB marker) were higher at 0 h time point (without IR exposure) in DSBR-deficient cells and remain much higher in DSBR-deficient compared to WT cells at 24 h after IR (Fig. 1d), confirming the association of DSBR-deficient status with EGFR-dependent miR-21 upregulation. However, we did notice that IR-stimulated EGFR and miR-21 levels remained up to 72 h in all examined cells including WT cells although IR-induced DSBs were repaired in WT cells at 24 h (Fig. 1d). This might be due to IR-induced DSBs activating multiple pathways that continually stimulate EGFR-dependent miR-21 upregulation indirectly even after IR-induced DSBs have been repaired. This is supported by our previous observation that increased miR-21 and EGFR levels were observed in the brains from WT mice at 1 year after (whole body) exposed to 0.5 Gy [32]. Of course, additional experiments are needed to elucidate the full mechanism.

b. Repair gene deficiency is the major reason for EGFR-dependent miR-21 upregulation in DSBR-deficient cells

To substantiate that increased miR-21 levels in DSBR-deficient cells were specifically due to DNA-PKcs or Rad54 deficiency, we knocked down DNA-PKcs or Rad54 in two different WT MEF lines from repeated experiments. When the DSBR gene (DNA-PKcs or Rad54) was knocked down in these WT MEFs, EGFR protein levels significantly increased (Fig. 2a, b, Supplementary Fig. 2) and the levels of miR-21 also increased (Fig. 2b). These results provide strong evidence to support that the increased EGFR-dependent miR-21 levels in DNA-PKcs−/− or Rad54−/− cells were due to a deficiency in the DSBR gene. Based on the results (similar phenotypes were observed in different WT MEFs), we performed the following experiments using only one WT MEF line (WT counterpart for Rad54−/− MEFs). We then examined the effects of STAT3 on miR-21 expression in these MEFs since STAT3 is a downstream effector of EGFR and regulates miR-21 expression [3]. Knocking down STAT3 in WT, DNA-PKcs−/− or Rad54−/− cells (Fig. 2c), significantly reduced miR-21 levels (Fig. 2d). Combining these data with the results shown in Fig. 1, we believe that DNA DSBs, even endogenous DSBs, are associated with stimulation of miR-21 expression. To confirm this, we performed the following experiments.

Most endogenous DNA DSBs occur during DNA replication, and are efficiently repaired by both NHEJ and HRR (more efficiently by HRR). To verify whether high levels of spontaneously generated DSBs are associated with stimulating EGFR-dependent miR-21 expression, we synchronized these cells and examined Egfr (mRNA) and miR-21 levels. The cells were synchronized in the S or G2/M phase using the approach described in Materials and methods (Fig. 3a). We counted co-localized (merged) γ-H2AX and 53BP1 foci as DSBs to avoid non-specific effects since it is known that γ-H2AX foci exist in the S phase cells without exogenous DNA damage stimulation. The results showed that DSB cells were more abundant during the S phase and less in the G2/M phase versus the asynchronous cells, particularly in DSBR-deficient Rad54−/− or DNA-PKcs−/− cells (Fig. 3b). Consistent with these data, Egfr (mRNA) and miR-21 levels were notably higher in the S-phase of Rad54−/− or DNA-PKcs−/− cells vs their WT counterparts, but there was no significant difference observed in the G2/M phase of these cells (Fig. 3c), further confirming the association of DSBs with EGFR-dependent miR-21 upregulation.

Fig. 3. Endogenous DSBs are responsible for upregulation of EGFR and miR-21 in DSBR-deficient cells.

Fig. 3.

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a. Cells were synchronized by treatment with nocodazole (G2/M phase) or double thymidine blockade (S-phase) as described in Materials and methods. Cell cycle distribution was measured by flow cytometry. NoSy, asynchronous cells. b. Upper panel: Representative immunostaining image for merged (co-localization) 53BP1 and γ-H2AX (scale bar = 30 μm) foci (DSB cells) in the S phase cells. Lower panel: Quantification of frequency of merged foci of 53BP1 and γ-H2AX (DSB cells) in non-synchronized (NoSy) and synchronized cells (S or G2/M). For each experiment, 50 cells per slide were counted. **, P<0.01; ***, P<0.001. c. Levels of Egfr mRNA and miR-21 in NoSy S or G2/M cells were analyzed by qPCR and were normalized to the value of NoSy WT cells. The graph depicts the mean and standard deviation from three independent experiments. N.S.D, no significant difference; *, P<0.05;***, P<0.01.

c. Inhibiting miR-21 expression suppresses the efficiency of IR-promoted cell growth in soft agar

Next, we were interested in investigating whether inhibiting miR-21 expression could suppress the efficiency of IR-promoted colony formation in soft agar. The MEFs used in this study were immortalized but these lines have a limited ability to form colonies in soft agar and are typically unable to form tumors in animals. We tested the efficiency of colony-formation in soft agar because acquisition of the capacity for anchorage-independent growth is a hallmark of carcinogenesis [16]. After these cells were exposed to 1 Gy, they were sub-cultured for 2 months. The cells were treated with control RNA or a miR-21 inhibitor (an RNA with the antisense sequence of miR-21). At 24 h after treatment, the cells were collected and divided into three portions to measure (i) miR-21 levels, (ii) plating efficiency, and (iii) soft agar colony formation efficiency. For experiments described in (ii) and (iii), additional fresh control RNA or a miR-21 inhibitor was added to medium (for anchorage-dependent growth) or soft agar (for anchorage-independent growth) until colony formation ended. We used non-irradiated miR-21 knock-in MEFs as a positive control. The miR-21 levels in DSBR-deficient cells were still much higher than in WT cells and the miR-21 inhibitor clearly reduced miR-21 level, although miR-21 levels were not eliminated by the inhibitor (Fig. 4a). The reason might be the miR-21 inhibitor acting as an antisense of miR-21, binding to the mature miR-21, which prevents binding to its targets, but may not completely degrade the miR-21; thus, qPCR could still detect some miR-21 signals. The cell plating efficiency at 2 months of subculture after IR exposure (Fig. 4b) was used to calibrate the soft agar colony-forming efficiency.

Fig. 4. Inhibition of miR-21 expression suppresses IR-promoted soft agar colony-forming efficiency.

Fig. 4.

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Non-irradiated miR-21 knock-in cells were used as a positive control. WT, Rad54−/− or DNA-PKcs−/− cells were exposed to 1 Gy, sub-cultured for 2 months, and treated with control RNA or a miR-21 inhibitor (20 nM). At 24 h after treatment, the cells were collected and divided into three portions to measure miR-21 level, anchorage-independent growth (plating efficiency) and anchorage-independent growth (soft agar colony formation) with freshly added control RNA or a miR-21 inhibitor. a. MiR-21 levels were measured using qPCR. The level of miR-21 from irradiated WT cells at 72 h after IR (Fig. 1a) is reference “1”. Data represent the mean and standard deviation from three independent experiments. **, P<0.01. b. Plating efficiency was assessed by colony forming ability (~ 1 week) from 200 cells in each dish containing 20 nM freshly added control RNA or a miR-21 inhibitor. These results were used to normalize the soft agar colony data. Data represent the mean and standard deviation from three independent experiments. c. Cells formed colonies ~ 3 weeks in soft agar (containing freshly added 20 Nm control RNA or a miR-21 inhibitor), the colonies were counted, the data were calibrated with the plating efficiency and then normalized with the value from miR-21 knock-in cells (defined as 100%). Data represent the mean and standard deviation for each cell line from three independent experiments. **, P<0.01; N.S.D, no significant difference.

MiR-21 knock-in MEFs produced 50–60 colonies from 2,000 cells, which is considered 100% for purposes of comparing to the oncogenic transformation propensity (soft agar colony-forming ability) of the other cell lines. IR (1 Gy) exposure significantly increased the soft agar colony-forming efficiency of WT and DSBR-deficient cells (Fig. 4c). Notably, the miR-21 inhibitor effectively decreased IR-induced soft agar colony-forming efficiency of all examined cells, more effectively decreased that of DNA-PKcs−/− or Rad54−/− cells, resulting in no significant difference in the efficiency between WT and the DSBR-deficient cells (Fig. 4c). These results suggest that the increased soft agar colony-forming efficiency in DSBR-deficient cells might be mainly due to increased EGFR-dependent miR-21.

To study whether other oncogenic miRNAs could also be associated with DSBs and elicit an effect similar to miR-21, we measured the miR-155 levels in WT, DNA-PKcs−/− or Rad54−/− cells with or without IR. MiR-155 is an oncogene that highly expressed in many types of cancers [33–36]. There was no difference in miR-155 levels between WT and the DSBR-deficient (DNA-PKcs−/− or Rad54−/−) cells with or without IR (Supplementary Fig. 3a). In addition, we also measured miR-34a levels in these cells since miR-34a, as a tumor suppressor, is a target of p53 and can be stimulated by IR [37, 38]. No apparent difference in miR-34a levels was found between WT and the DSBR-deficient cells (Supplementary Fig. 3b). These results provide additional evidence that EFDR-dependent miR-21 is associated with DSB-induced genomic instability, although we cannot completely exclude other miRNA involvements.

d. Activated ATM and ATR may also contribute to IR-induced miR-21 upregulation

We reported previously that IR-stimulated miR-21 upregulation was due to the activation of AP-1 and EGFR further activating STAT3 [3]. Recently, it was found that DNA DSB-activated ATM and ATR contributed to STAT3 activation [39]. These results suggest that both ATM and ATR may be involved in IR-stimulated miR-21 expression. To test this possibility, we examined miR-21 levels in WT, DNA-PKcs−/− or Rad54−/− cells using a specific ATM or ATR inhibitor. We irradiated cells with 5 Gy to enhance the phosphorylation of ATM and ATR. Without IR, the ATM or ATR inhibitor did not significantly affect the miR-21 levels in these MEFs (data not shown). Although inhibition of ATM or ATR alone did not significantly reduce IR-stimulated miR-21 level in all examined cells including WT and DSBR-deficient cells (Supplementary Fig. 4a), combined inhibition of both ATM and ATR significantly reduced IR-stimulated miR-21 levels in WT cells, and more significantly in DNA-PKcs−/− or Rad54−/− cells (Supplementary Fig. 4a). These results suggest that ATM/ATR may also (mildly) contribute to DSB-promoted miR-21 upregulation.

To further confirm the effects of ATM/ATR on IR-stimulated miR-21 expression, we compared the effects of an ATM, ATR or EGFR inhibitor on IR-induced EGFR phosphorylation, STAT3 phosphorylation and c-Jun (an AP-1 subunit) level in WT and the DSBR-deficient cells since AP-1 is another key factor in modulating miR-21 expression, and EGFR, ATM or ATR could be directly or indirectly involved in AP-1 expression/activation [40–44]. Interestingly, although EGFR inhibition did not affect ATM/ATR phosphorylation (data not shown), ATM/ATR inhibition significantly decreased EGFR phosphorylation (Supplementary Fig. 4b), supporting that ATM/ATR might be associated with miR-21 upregulation through stimulating EGFR activity. The STAT3 phosphorylation status provided additional evidence to support this (Supplementary Fig. 4c). Furthermore, ATM/ATR inhibition also decreased c-Jun levels in these irradiated cells (Supplementary Fig. 4d), supporting the association of ATM/ATR with IR-stimulated miR-21 upregulation. However, we did not observe any significant difference in the effects of ATM/ATR inhibition on these phosphorylated or protein levels between irradiated WT and the DSBR-deficient cells (Supplementary Fig. 4b-d) as shown in miR-21 levels (Supplementary Fig. 4a). This might be explained by that the sensitivity limitations of western blot is not easy to detect ATM/ATR-induced mild effects on phosphorylation or protein levels, which is different from the more sensitive PCR assay used to detect changes in miR-21 level. Based on the results shown in this study, we developed a model to define the possible relationship between DNA DSBs, upregulation of miR-21 and tumorigenesis (Fig. 5). This model hints that (i) DSBs are associated with EGFR-dependent mR-21 upregulation; (ii) mR-21 upregulation is associated with promotion of oncogenic transformation and further tumorigenesis; (iii) IR-activated ATM/ATR-promoted STAT3 and AP-1 may stimulate miR-21 expression in a direct or indirect way.

Fig. 5. Schematic illustration of the possible relationship between DSBs, miR-21 and oncogenic transformation.

Fig. 5.

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DSBs stimulate upregulation of EGFR, which is also mildly affected by IR-activated of ATM/ATR through their substrates. EGFR activates STAT3 that in turn stimulates miR-21 upregulation. IR-stimulated AP-1 directly stimulates miR-21 expression, and EGFR, ATM or ATR may also stimulate IR-induced AP-1. Upregulated miR-21 might play an important role in promoting oncogenic transformation propensity and subsequent tumorigenesis.

4. Discussion and conclusions

In this study, we focused on elucidating the relationship between DSBR, EGFR-dependent miR-21 expression and IR-promoted soft agar colony-formation efficiency. Our data suggest that cells deficient in the HRR or NHEJ pathway have increased the efficiency of IR-induced cell growth in soft agar, which is associated with EGFR-dependent miR-21 upregulation.

Previously, it was reported that miR-21 upregulation resulted in cell resistance to IR [45]. We also showed that miR-21 stimulates DNA DSBR by promoting both NHEJ and HRR pathways through targeting GSK3B [46]. However, the higher miR-21 levels in Rad54- or DNA-PKcs-deficient cells does not result in cell resistance to IR, since these cells are deficient in DSBR. Blocking miR-21 function (even though DSBs are still present in the cells and continually associated with EGFR-dependent miR-21 expression), results in a significant decrease in soft agar colony formation efficiency. These results suggest that miR-21 upregulation is associated with DSB-promoted cell growth efficiency in soft agar. Before we reported that IR-activated EGFR plays a major role in promoting cell colony forming efficiency in soft agar [30], and the results shown in this study support miR-21 serving as a major downstream effector of EGFR for such efficiency. Whereas, we cannot exclude the effects of other downstream effectors of EGFR or other gene mutations on this process, since blocking miR-21 in irradiated cells still resulted in higher efficiency of cell growth in soft agar versus non-irradiated cells.

ATM and ATR have dual effects on DNA damage-induced tumorigenesis. On one hand, ATM and ATR serving as the main DNA damage response mediators, play an important role in maintaining genomic integrity [47–49]. On the other hand, DNA damage-induced ATM/ATR over-activation promotes tumorigenesis [39], which is associated with the activation of their downstream pathways. As described in this study, IR-activated ATM/ATR might also promote miR-21 expression, which involves EGFR/STAT3 and/or AP-1 activation, the underlying mechanism requires further study to elucidate. These results suggest that miR-21 upregulation may be a downstream effector of IR-induced ATM/ATR over-activation on tumorigenesis. The opposite effects of ATM/ATR on genomic integrity may depend on the cell’s response to acute or chronic DNA damage. Acute DNA damage (like single high dose)-activated ATM/ATR plays a protective role in maintaining genomic integrity through promoting DNA repair [50]. However, chronic DNA damage (such as persistent existence of DNA DSB-activated ATM/ATR), might play a role in promoting genomic instability through activating EGFR/STAT3 and/or AP-1-stimulated miR-21. Furthermore, the generation of small non-coding RNA at DSB sites in mammalian cells may directly or indirectly affect DSBR as well [51–53]. Therefore, the DSB-dependent generation of non-coding RNA and the concomitant cellular response may also influence miR-21 regulation and DSB-promoted cell growth in soft agar, which warrants further study in the near future.

In this study, we observed that miR-21 inhibition significantly suppressed cell growth in soft agar of irradiated cells, suggesting that miR-21 is an important factor associated with IR-induced soft agar colony-forming efficiency. Notably, miR-21 inhibition eliminated the difference in soft agar colony-forming efficiency between WT and DSBR-deficient cells, supporting that DSB-associated miR-21 upregulation contributes to transformed cell growth. Taken together, our results show for the first time that DSBR deficiency-stimulated cell growth in soft agar is associated with EGFR-dependent miR-21 upregulation. These findings contribute towards our understanding of the mechanisms, by which the DSBR system-involved regulation of miRNA expression protects genome integrity and prevents tumorigenesis.

Supplementary Material

1

Highlights.

  • DNA double strand breaks are associated with EGFR-dependent miR-21 up-regulation.

  • Over-activated ATM and ATR may also contribute to ionizing radiation-induced miR-21 up-regulation.

  • MiR-21 up-regulation is associated with ionizing radiation-promoted cell growth in soft agar.

Acknowledgements

We thank Drs. Kevin Mills, Gloria Li, David Chen, Chuan-Yuan Li and George Iliakis for providing cell lines; as well as Dr. William Dynan and Ms. Doreen Theune for editing the manuscript. This work was supported by grants from US National Institutes of Health (CA186129 and CA185882 to Y.W., and P30CA138292 to Winship Cancer Institute), and the National Aeronautics and Space Administration, (NNX11AC30G to Y.W.).

Abbreviations used:

IR

ionizing radiation

miRNA

microRNA

DSB

double strand break

SSB

single strand break

DSBR

double strand break repair

SSBR

single strand break repair

NHEJ

non-homologous end-joining

HRR

homologous recombination repair

MEFs

murine embryonic fibroblasts

miRNA

microRNA

ctRNA

control RNA

DNA-PKcs

DNA protein kinase catalytic subunit

WT

wild-type

qPCR

quantitative real-time PCR

siRNA

small interfering RNA

Footnotes

Conflict of interest

The authors have no conflict of interest to declare with regard to the content of this manuscript.

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References

  • 1.Volinia S, Calin GA, Liu C-G, Ambs S, Cimmino A, Petrocca F, Visone R, Iorio M, Roldo C, Ferracin M, Prueitt RL, Yanaihara N, Lanza G, Scarpa A, Vecchione A, Negrini M, Harris CC, Croce CM. A microRNA expression signature of human solid tumors defines cancer gene targets. Proceedings of the National Academy of Sciences of the United States of America. 2006;103:2257–61. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Croce CM. miRNAs in the spotlight: Understanding cancer gene dependency. Nat Med. 2011;17:935–6. [DOI] [PubMed] [Google Scholar]
  • 3.Zhu Y, Yu X, Fu H, Wang H, Wang P, Zheng X, Wang Y. MicroRNA-21 involves radiation-promoted liver carcinogenesis. Int J Clin Exp Med. 2010;3:211–22. [PMC free article] [PubMed] [Google Scholar]
  • 4.Medina PP, Nolde M, Slack FJ. OncomiR addiction in an in vivo model of microRNA-21induced pre-B-cell lymphoma. Nature. 2010;467:86–90. [DOI] [PubMed] [Google Scholar]
  • 5.Hatley ME, Patrick DM, Garcia MR, Richardson JA, Bassel-Duby R, van Rooij E, Olson EN. Modulation of K-Ras-Dependent Lung Tumorigenesis by MicroRNA-21. Cancer Cell. 2010;18:282–93. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Feng Y-H, Tsao C-J. Emerging role of microRNA-21 in cancer. Biomedical reports. 2016;5:395–402. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Ward JF. Biochemistry of DNA lesions. Radiat Res. 1985;104:S103–S11. [PubMed] [Google Scholar]
  • 8.Cheng Y, Liu X, Zhang S, Lin Y, Yang J, Zhang C. MicroRNA-21 protects against the H2O2-induced injury on cardiac myocytes via its target gene PDCD4. Journal of Molecular and Cellular Cardiology. 2009;47:5–14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Lin Y, Liu X, Cheng Y, Yang J, Huo Y, Zhang C. Involvement of MicroRNAs in hydrogen peroxide-mediated gene regulation and cellular injury response in vascular smooth muscle cells. J Biol Chem. 2009;284:7903–13. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Syed DN, Khan MI, Shabbir M, Mukhtar H. MicroRNAs in skin response to UV radiation. Current drug targets. 2013;14:1128–34. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Kurimasa A, Ouyang H, Dong L-j, Wang S, Li X, Cordon-Cardo C, Chen DJ, Li GC. Catalytic subunit of DNA-dependent protein kinase: Impact on lymphocyte development and tumorigenesis. Proceedings of the National Academy of Sciences, USA. 1999;96:1403–8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Essers J, Hendriks RW, Swagemakers SMA, Troelstra C, de Wit J, Bootsma D, Hoeijmakers JHJ, Kanaar R. Disruption of Mouse RAD54 Reduces Ionizing Radiation Resistance and Homologous Recombination. Cell. 1997;89:195–204. [DOI] [PubMed] [Google Scholar]
  • 13.Fisher AEO, Hochegger H, Takeda S, Caldecott KW. Poly(ADP-Ribose) Polymerase 1 Accelerates Single-Strand Break Repair in Concert with Poly(ADP-Ribose) Glycohydrolase. Mol Cell Biol. 2007;27:5597–605. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Hakem R. DNA-damage repair; the good, the bad, and the ugly. EMBO J. 2008;27:589–605. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Polo LM, Xu Y, Hornyak P, Garces F, Zeng Z, Hailstone R, Matthews SJ, Caldecott KW, Oliver AW, Pearl LH. Efficient Single-Strand Break Repair Requires Binding to Both Poly(ADP-Ribose) and DNA by the Central BRCT Domain of XRCC1. Cell Reports. 2019;26:573–81.e5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Borowicz S, Van Scoyk M, Avasarala S, Karuppusamy Rathinam MK, Tauler J, Bikkavilli RK, Winn RA. The soft agar colony formation assay. Journal of visualized experiments : JoVE. 2014:e51998-e. [DOI] [PMC free article] [PubMed]
  • 17.Mills KD, Ferguson DO, Essers J, Eckersdorff M, Kanaar R, Alt FW. Rad54 and DNA Ligase IV cooperate to maintain mammalian chromatid stability. Genes Dev. 2004;18:1283–92. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Liu X, He Y, Li F, Huang Q, Kato Takamitsu A, Hall Russell P, Li C-Y. Caspase-3 Promotes Genetic Instability and Carcinogenesis. Mol Cell. 2015;58:284–96. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Wachsberger PR, Li WH, Guo M, Chen D, Cheong N, Ling CC, Li G, Iliakis G. Rejoining of DNA Double-Strand Breaks in Ku80-Deficient Mouse Fibroblasts. Radiat Res. 1999;151:398–407. [PubMed] [Google Scholar]
  • 20.Wang J, Zhang X, Wang P, Wang X, Farris A, Wang Y. Lessons learned using different mouse models during space radiation-induced lung tumorigenesis experiments. Life Sci Space Res. 2016;9:48–55. [DOI] [PubMed] [Google Scholar]
  • 21.Tang S, Liu B, Liu J, Wang J, Wang Y. A protein-mRNA feedback exists in miR-21-associated E-Selectin expression Int J Radiat Biol. 2019;In Press. [DOI] [PMC free article] [PubMed]
  • 22.Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 2001;25:402–8. [DOI] [PubMed] [Google Scholar]
  • 23.Wang HY, Liu S, Zhang P, Zhang S, Naidu M, Wang HC, Wang Y. S-phase cells are more sensitive to high linear energy transfer radiation. Int J Rad Onc Bio Phys. 2009;74:1236–41. [DOI] [PubMed] [Google Scholar]
  • 24.Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM. DNA Double-stranded Breaks Induce Histone H2AX Phosphorylation on Serine 139. J Biol Chem. 1998;273:5858–68. [DOI] [PubMed] [Google Scholar]
  • 25.Wang B, Matsuoka S, Carpenter PB, Elledge SJ. 53BP1, a Mediator of the DNA Damage Checkpoint. Science. 2002;298:1435–8. [DOI] [PubMed] [Google Scholar]
  • 26.Rothkamm K, Lobrich M. Evidence for a lack of DNA double-strand break repair in human cells exposed to very low x-ray doses. Proc Natl Acad Sci USA 2003;100:5057–62. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Das BB, Huang S-yN, Murai J, Rehman I, Amé J-C, Sengupta S, Das SK, Majumdar P, Zhang H, Biard D, Majumder HK, Schreiber V, Pommier Y. PARP1–TDP1 coupling for the repair of topoisomerase I–induced DNA damage. Nucleic Acids Res. 2014;42:4435–49. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Zhang X, Ng W-L, Wang P, Tian L, Werner E, Wang H, Doetsch P, Wang Y. MicroRNA-21 Modulates the Levels of Reactive Oxygen Species by Targeting SOD3 and TNFα. Cancer Res. 2012;72:4707–13. [DOI] [PMC free article] [PubMed] [Google Scholar] [Research Misconduct Found]
  • 29.Ng WL, Chen G, Wang M, Wang H, Story M, Shay JW, Zhang X, Wang J, Amin ARMR, Hu B, Cucinotta FA, Wang Y. OCT4 as a target of miR-34a stimulates p63 but inhibits p53 to promote human cell transformation. Cell Death Dis. 2014;5:e1024. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  • 30.Wang J, Farris III A, Xu K, Wang P, Zhang X, Duong D, Yi H, Shu H-K, Sun S-Y, Wang Y. GPRC5A suppresses protein synthesis at the endoplasmic reticulum to prevent radiation-promoted lung tumorigenesis. Nature Communications. 2016;7:11795. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Wei L, Nakajima S, Hsieh CL, Kanno S, Masutani M, Levine AS, Yasui A, Lan L. Damage response of XRCC1 at sites of DNA single strand breaks is regulated by phosphorylation and ubiquitylation after degradation of poly(ADP-ribose). J Cell Sci. 2013;126:4414–23. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Shi Y, Zhang X, Tang X, Wang P, Wang H, Wang Y. MiR-21 Is Continually Elevated Long-Term in the Brain following Ionizing Radiation. Radiat Res. 2012;177:124–8. [DOI] [PubMed] [Google Scholar]
  • 33.Faraoni I, Antonetti FR, Cardone J, Bonmassar E. miR-155 gene: A typical multifunctional microRNA. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease. 2009;1792:497–505. [DOI] [PubMed] [Google Scholar]
  • 34.Donnem T, Eklo K, Berg T, Sorbye SW, Lonvik K, Al-Saad S, Al-Shibli K, Andersen S, Stenvold H, Bremnes RM, Busund LT. Prognostic impact of MiR-155 in non-small cell lung cancer evaluated by in situ hybridization. J Transl Med. 2011;9:6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Mattiske S, Suetani RJ, Neilsen PM, Callen DF. The oncogenic role of miR-155 in breast cancer. Cancer Epidemiol Biomarkers Prev. 2012;21:1236–43. [DOI] [PubMed] [Google Scholar]
  • 36.Xue X, Liu Y, Wang Y, Meng M, Wang K, Zang X, Zhao S, Sun X, Cui L, Pan L, Liu S. MiR-21 and MiR-155 promote non-small cell lung cancer progression by downregulating SOCS1, SOCS6, and PTEN. Oncotarget. 2016;7:84508–19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Chang T, Wentzel E, Kent O, Ramachandran K, Mullendore M, Lee K, Feldmann G, Yamakuchi M, Ferlito M, Lowenstein C, Arking D, Beer M, Maitra A, Mendell J. Trans-activation of miR-34a by p53 broadly influences gene expression and promotes apoptosis. Mol Cell. 2007;26:745–52. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.He L, He X, Lim LP, de Stanchina E, Xuan Z, Liang Y, Xue W, Zender L, Magnus J, Ridzon D, Jackson AL, Linsley PS, Chen C, Lowe SW, Cleary MA, Hannon GJ. A microRNA component of the p53 tumour suppressor network. Nature. 2007;447:1130–4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Liu X, Li F, Huang Q, Zhang Z, Zhou L, Deng Y, Zhou M, Fleenor DE, Wang H, Kastan MB, Li C-Y. Self-inflicted DNA double-strand breaks sustain tumorigenicity and stemness of cancer cells. Cell Res. 2017;27:764–83. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Barilá D, Mangano R, Gonfloni S, Kretzschmar J, Moro M, Bohmann D, Superti-Furga G. A nuclear tyrosine phosphorylation circuit: c-Jun as an activator and substrate of c-Abl and JNK. EMBO J. 2000;19:273–81. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Li G, Gustafson-Brown C, Hanks SK, Nason K, Arbeit JM, Pogliano K, Wisdom RM, Johnson RS. c-Jun is essential for organization of the epidermal leading edge. Dev Cell. 2003;4:865–77. [DOI] [PubMed] [Google Scholar]
  • 42.Foray N, Marot D, Gabriel A, Randrianarison V, Carr AM, Perricaudet M, Ashworth A, Jeggo P. A subset of ATM- and ATR-dependent phosphorylation events requires the BRCA1 protein. EMBO J. 2003;22:2860–71. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Kurz EU, Lees-Miller SP. DNA damage-induced activation of ATM and ATM-dependent signaling pathways. DNA Repair (Amst). 2004;3:889–900. [DOI] [PubMed] [Google Scholar]
  • 44.Normanno N, De Luca A, Bianco C, Strizzi L, Mancino M, Maiello MR, Carotenuto A, De Feo G, Caponigro F, Salomon DS. Epidermal growth factor receptor (EGFR) signaling in cancer. Gene. 2006;366:2–16. [DOI] [PubMed] [Google Scholar]
  • 45.Cho B, Kim H, Lee D, Choi E, Hwang Y, Chun S, Kim I. MicroRNA-21 inhibitor potentiates anti-tumor effect of radiation therapy in vitro and in vivo. Tumor Microenviroment. 2014;2:1–13. [Google Scholar]
  • 46.Hu B, Wang X, Hu S, Ying X, Wang P, Zhang X, Wang J, Wang H, Wang Y. miR-21-mediated Radioresistance Occurs via Promoting Repair of DNA Double Strand Breaks. J Biol Chem. 2017;292:3531–40. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  • 47.Bartkova J, Horejsi Z, Koed K, Kramer A, Tort F, Zieger K, Guldberg P, Sehested M, Nesland JM, Lukas C, Orntoft T, Lukas J, Bartek J. DNA damage response as a candidate anti-cancer barrier in early human tumorigenesis. Nature. 2005;434:864–70. [DOI] [PubMed] [Google Scholar]
  • 48.Gorgoulis VG, Vassiliou L-VF, Karakaidos P, Zacharatos P, Kotsinas A, Liloglou T, Venere M, DiTullio RA Jr, Kastrinakis NG, Levy B, Kletsas D, Yoneta A, Herlyn M, Kittas C, Halazonetis TD. Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions. Nature. 2005;434:907–13. [DOI] [PubMed] [Google Scholar]
  • 49.Marechal A, Zou L. DNA damage sensing by the ATM and ATR kinases. Cold Spring Harb Perspect Biol. 2013;5. [DOI] [PMC free article] [PubMed]
  • 50.Golding SE, Rosenberg E, Khalil A, McEwen A, Holmes M, Neill S, Povirk LF, Valerie K. Double strand break repair by homologous recombination is regulated by cell cycle-independent signaling via ATM in human glioma cells. J Biol Chem. 2004;279:15402–10. [DOI] [PubMed] [Google Scholar]
  • 51.Wei W, Ba Z, Gao M, Wu Y, Ma Y, Amiard S, White Charles I, Rendtlew Danielsen Jannie M, Yang Y-G, Qi Y. A Role for Small RNAs in DNA Double-Strand Break Repair. Cell. 2012;149:101–12. [DOI] [PubMed] [Google Scholar]
  • 52.Francia S, Michelini F, Saxena A, Tang D, de Hoon M, Anelli V, Mione M, Carninci P, d/’Adda di Fagagna F. Site-specific DICER and DROSHA RNA products control the DNA-damage response. Nature. 2012;488:231–5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Liu B, Liu M, Wang J, Zhang X, Wang X, Wang P, Wang H, Li W, Wang Y. DICER dependent biogenesis of let-7 miRNAs affects human cell response to DNA damage via targeting p21/p27 Nucleic Acids Res. 2015;43:1626–36. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]

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