Research ArticleCancer

IDH1-R132H acts as a tumor suppressor in glioma via epigenetic up-regulation of the DNA damage response

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Science Translational Medicine  13 Feb 2019:
Vol. 11, Issue 479, eaaq1427
DOI: 10.1126/scitranslmed.aaq1427

Linking glioma metabolism and DNA repair

Mutations in isocitrate dehydrogenase 1 (IDH1) are frequently found in gliomas and are associated with better outcomes. Núñez et al. discovered that, in addition to its roles in metabolism and epigenetics, mutant IDH1 also helps maintain genomic stability in tumors by enhancing the DNA damage response. This finding helps explain why patients with IDH1-mutant tumors have better survival despite their tumors being less sensitive to radiation than other gliomas. The authors also examined the mechanism for this phenomenon in mouse models and demonstrated that pharmacological inhibition of the DNA damage response sensitizes IDH1-mutant tumors to radiation, suggesting a potential direction for further therapeutic advances.

Abstract

Patients with glioma whose tumors carry a mutation in isocitrate dehydrogenase 1 (IDH1R132H) are younger at diagnosis and live longer. IDH1 mutations co-occur with other molecular lesions, such as 1p/19q codeletion, inactivating mutations in the tumor suppressor protein 53 (TP53) gene, and loss-of-function mutations in alpha thalassemia/mental retardation syndrome X-linked gene (ATRX). All adult low-grade gliomas (LGGs) harboring ATRX loss also express the IDH1R132H mutation. The current molecular classification of LGGs is based, partly, on the distribution of these mutations. We developed a genetically engineered mouse model harboring IDH1R132H, TP53 and ATRX inactivating mutations, and activated NRAS G12V. Previously, we established that ATRX deficiency, in the context of wild-type IDH1, induces genomic instability, impairs nonhomologous end-joining DNA repair, and increases sensitivity to DNA-damaging therapies. In this study, using our mouse model and primary patient-derived glioma cultures with IDH1 mutations, we investigated the function of IDH1R132H in the context of TP53 and ATRX loss. We discovered that IDH1R132H expression in the genetic context of ATRX and TP53 gene inactivation (i) increases median survival in the absence of treatment, (ii) enhances DNA damage response (DDR) via epigenetic up-regulation of the ataxia-telangiectasia–mutated (ATM) signaling pathway, and (iii) elicits tumor radioresistance. Accordingly, pharmacological inhibition of ATM or checkpoint kinases 1 and 2, essential kinases in the DDR, restored the tumors’ radiosensitivity. Translation of these findings to patients with IDH1132H glioma harboring TP53 and ATRX loss could improve the therapeutic efficacy of radiotherapy and, consequently, patient survival.

INTRODUCTION

Mutated isocitrate dehydrogenase 1 (IDH1R132H) is found in 80% of low-grade gliomas (LGGs) [World Health Organization (WHO) grade II or III] and in a subset of high-grade gliomas (WHO grade IV) (13). Two main molecular subtypes of glioma, which harbor IDH1R132H, express the following: (i) IDH1R132H, 1p/19q codeletion, and TERT promoter mutations; or (ii) IDH1R132H, mutant TP53, and inactivation of ATRX (35). In spite of a better prognosis, 50 to 75% of IDH1R132H gliomas undergo malignant transformation over time, becoming WHO grade IV glioblastomas (GBMs) (2, 6).

IDH1R132H has been identified as an early event in glioma development, preceding TP53 and ATRX mutations (7, 8). IDH1R132H is a gain-of-function mutation that converts α-ketoglutarate to (R)-2-hydroxyglutarate (2HG) (911). 2HG inhibits DNA and histone demethylases, namely the ten-eleven translocation enzymes and lysine demethylases, respectively, resulting in hypermethylation of DNA and histones (10, 11). This elicits epigenetic reprogramming of the IDH1R132H tumor cells’ transcriptome (1013). However, the molecular mechanisms that mediate increased survival in patients with mutant IDH1 (mIDH1) glioma remain unknown.

Genomic instability is prevalent in gliomas; it is thought to promote tumorigenesis and an aggressive phenotype (4, 14). DNA damage response (DDR) maintains genomic stability, senses DNA damage, and regulates the mitotic cell cycle progression and DNA repair mechanisms (15). Ataxia-telangiectasia–mutated (ATM), a member of phosphatidylinositol 3-kinase (PI3K)–like protein kinase family, plays a critical role in these processes (16).

Here, we demonstrate that IDH1R132H, in the context of ATRX and TP53 knockdown (KD), increases DDR activity, enhancing genomic stability and extending median survival (MS) in our mIDH1 mouse glioma model. We demonstrate that 2HG induces hypermethylation of histone 3 (H3), which elicits epigenetic reprogramming of the tumor cells’ trancriptome. RNA sequencing (RNA-seq), bromouridine sequencing (Bru-seq), and chromatin immunoprecipitation–sequencing (ChIP-seq) data from mIDH1 tumors uncovered enrichment of gene ontologies (GOs) related to DDR, genomic stability, and activation of DNA repair pathways, such as ATM signaling and homologous recombination DNA repair (HR repair). Consequently, mIDH1 tumors exhibited enhanced DDR. Increases in DDR activity were observed in mIDH1 human glioma cells from surgical biopsies. In addition, radiation failed to increase survival in mIDH1 tumor–bearing animals. Pharmacological inhibition of DDR conferred radiosensitivity in mIDH1 tumor–bearing mice, resulting in prolonged MS. Our findings indicate that DDR inhibition in combination with radiation could provide a therapeutic strategy for patients with IDH1R132H glioma harboring ATRX and TP53 inactivating mutations.

RESULTS

mIDH1 mouse glioma model exhibits increased survival and inhibition of oligodendrocyte differentiation

We generated a mIDH1 mouse glioma model using the Sleeping Beauty (SB) transposon system (4, 17) to uncover the impact of IDH1R132H in the context of ATRX and TP53 loss. Gliomas were induced by receptor tyrosine kinase (RTK)/RAS/PI3K activation in combination with shp53, shATRX, and IDH1R132H (fig. S1A). Mice from the three experimental groups, namely, (i) control (NRAS G12V-shp53), (ii) wild-type (wt) IDH1 (NRAS G12V-shp53-shATRX), and (iii) mIDH1 (NRAS G12V-shp53-shATRX-IDH1R132H), developed brain tumors (Fig. 1A and fig. S1B). The most aggressive tumor was wt-IDH1 (MS = 70 days). Notably, IDH1R132H increased MS (163 days; P < 0.0001) (Fig. 1A). In all groups, tumor cells did not express myosin VIIa (fig. S1, C and D), indicating that they did not originate from cells in the ependymal layer of the lateral ventricle. Because of the use of the shATRX construct to generate the wt-IDH1 and mIDH1 tumor models, ATRX expression was suppressed in these tumors (fig. S1E). IDH1R132H expression was only positive in mIDH1 tumors (fig. S1F). Wt-IDH1 and mIDH1 tumors (fig. S1G) expressed phosphorylated extracellular signal–regulated kinases 1 and 2, consistent with RTK activation observed in human mIDH1 and wt-IDH1 gliomas (fig. S1, H to K). We generated neurospheres (NSs) from mouse glioma subgroups (fig. S2A). Both wt-IDH1–NS and mIDH1-NS exhibited alternative lengthening of telomeres (ALT), which was associated with the presence of shATRX, whereas ALT was not detected in control NS or normal mouse brain (fig. S2B). IDH1R132H expression was confirmed in mIDH1-NS (Fig. 1B), in human glioma cells stably transfected with IDH1R132H (fig. S2C), and in human glioma cells with endogenous expression of IDH1R132H, TP53, and ATRX inactivating mutations (fig. S2D). In mIDH1-NS, 2HG concentration was, on average, 8.16 μg/mg of protein (Fig. 1C). We observed a reduction in 2HG production in mIDH1-NS (~4-fold; P < 0.0001) after treatment with AGI-5198, an IDH1R132H inhibitor, equivalent to the basal amount of wt-IDH1–NS (Fig. 1C). AGI-5198 inhibited cell viability (fig. S2E) and proliferation (2.8-fold; P < 0.0001) (fig. S2F) in mIDH1-NS, consistent with previous results in human glioma cells (18). Earlier reports indicated that IDH1R132H suppresses cellular differentiation (13, 18); thus, we evaluated the expression of oligodendrocyte and astrocyte differentiation markers. RNA-seq analysis revealed that a group of differentially expressed (DE) genes, involved in cell differentiation pathways, were down-regulated in mIDH1 tumors (Fig. 1D and fig. S3A). Gene set enrichment analysis (GSEA) (Fig. 1E and fig. S3) suggests that IDH1R132H inhibits differentiation in our model. Down-regulated GO terms in mIDH1-NS included Olig2 and Mbp (Fig. 1, E to G). mIDH1 tumors exhibited decreased amounts of (i) oligodendrocyte transcription factor 2 (OLIG2) (2.3-fold; P < 0.05), (ii) myelin basic protein (MBP) (9.3-fold; P < 0.001), and (iii) glial fibrillary acidic protein (GFAP) (6.7-fold; P < 0.05) (fig. S4A). In addition, mIDH1 tumors exhibited increased SOX2 [SRY (sex determining region Y)–box 2] expression (fig. S4B) and no change in PAX3 (paired box 3) expression (fig. S4C). In agreement with the in vivo data, wt-IDH1–NS expressed more OLIG2 than mIDH1-NS (fig. S4D). Inhibition of mIDH1 using AGI-5198 did not affect OLIG2 expression in wt-IDH1–NS, whereas it induced OLIG2 expression in mIDH1-NS (fig. S4D). The impact of mIDH1 on cell differentiation was confirmed in mIDH1-NS by immunofluorescence for OLIG2, GFAP, and nestin, markers for oligodendrocytes, astrocytes, and undifferentiated neural progenitor cells, respectively. Treatment of mIDH1-NS with AGI-5198 and retinoic acid enhanced OLIG2 and GFAP expression (fig. S4E).

Fig. 1 IDH1R132H increases MS and inhibits cell differentiation in a mouse glioma model.

(A) Kaplan-Meier survival curves for mice bearing mIDH1 (n = 24), wt-IDH1 (n = 10), or control (n = 12) gliomas (P < 0.0001, Mantel-Cox test). (B) Western blot (WB) analysis using NS from control (Ctrl), wt-IDH1, and mIDH1 tumors. Data are shown as IDH1R132H and total IDH1 expression. β-Actin, loading controls. (C) 2HG expression in mouse NS in the presence or absence of 1.5 μM AGI-5198. ****P < 0.0001, unpaired t test; n.s., not significant. Data are shown as means ± SEM (n = 3 biological replicates). (D) Differential gene expression in mIDH1 tumors analyzed by RNA-seq. Volcano plot showing DE genes in mIDH1 versus wt-IDH1 mouse NS. The −log10 q values were plotted against the log2 fold change in gene expression. Up-regulated genes [n = 906; ≥1.5-fold; false discovery rate (FDR)–corrected P < 0.05] are depicted as red dots; genes that were down-regulated (n = 1067; ≥1.5-fold; FDR-corrected P < 0.05) are depicted in green. Orange triangles represent down-regulated genes involved in oligodendrocyte differentiation. Blue triangles represent up-regulated genes involved in DDR. FDR-corrected P values = q values; two-sided moderated Student’s t test (n = 3 biological replicates per group). (E to G) Pathway enrichment maps of DE genes in mIDH1-NS versus wt-IDH1–NS. Clusters of nodes depicted in green (E) illustrate differentially down-regulated pathways resulting from GSEA (P < 0.05; overlap cutoff, >0.5) (see full map in fig. S3). The yellow highlighted nodes indicate down-regulated GO terms containing Olig2 (F) and Mbp (G).

We also investigated the tumor-initiating stem cell frequency using a limiting dilution assay (LDA). The LDA assay results indicate that both mouse NS and human mIDH1 glioma cells have lower stem cell frequency than wt cells (fig. S5, A to J). In vivo analysis for tumor-initiating cells (TICs) showed that 100% of animals generated tumors and succumbed because of tumor burden after implantation of 30 × 105, 10 × 105, 3 × 103, and 1 × 103 wt-IDH1 cells, whereas with mIDH1 cells, only 40% of animals generated tumors when implanted with 1 × 103 cells (fig. S5, K and L). These results suggest that there is a lower number of TICs among mIDH1 than among wt-IDH1 glioma cells. We also analyzed differences in the cell cycle profiles in our model in vivo. Frequencies of mitotic (pH3Ser10+) and actively proliferating (EdU+) glioma cells were higher in wt-IDH1 tumors (P < 0.0001; fig. S6, A to F).

IDH1R132H induces H3 hypermethylation at genomic regions associated with DDR pathways

WB analysis in mIDH1-NS demonstrated increased amounts of H3K4me3 (1.9-fold; P < 0.01), H3K27me3 (2.3-fold; P < 0.01), and H3K36me3 (7.1-fold; P < 0.01) (Fig. 2A). Similar amounts of H3K4me1 (Fig. 2A) and H3K79me2 (fig. S7, A and B) marks were observed in all tumor genotypes. mIDH1 tumor sections exhibited increased amounts of H3K27me3 (4.6-fold; P < 0.001) and H3K36me3 (7.3-fold; P < 0.001) (fig. S7C). There were no changes in mIDH1-NS H3 acetylation (fig. S7, D and E). H3 hypermethylation was also observed in human glioma cells harboring IDH1R132H (fig. S7F).

Fig. 2 IDH1R132H increases histone hypermethylation and elicits epigenetic enrichment of GOs related to DDR.

(A) WB assay performed on control (Ctrl), wt-IDH1–NS (wt), and mIDH1-NS (mut) for H3K4me3, H3K27me3, H3K36me3, and H3K4me1 marks. Total histone H3, loading control. The bar graph represents the semi-quantification of the histone bands (n = 3 technical replicates). **P < 0.01; ***P < 0.001; one-way analysis of variance (ANOVA). (B and C) ChIP-seq enrichment across the genome shows differential peaks of histone marks in mIDH1-NS. Heat maps show H3K4me3 (B) and H3K27me3 (C) peaks ±2 kilo–base pair (kbp), with each row representing a distinct peak. The blue-to-red color gradient indicates high-to-low counts in the corresponding regions. The heat maps show three biological replicates per group for wt-IDH1–NS and mIDH1-NS. (D to H) Distribution of histone marks within specific genomic regions. (D) Diagram represents known genome annotations. CDS, coding sequence. (E and F) Bar graphs represent the specific genomic regions where H3K4me3 is enriched in wt-IDH1–NS (E) and mIDH1-NS (F). (G and H) Bar graphs represent the specific genomic regions where H3K27me3 is enriched in wt-IDH1–NS (G) and mIDH1-NS (H). Red bars show ChIP-seq data, and gray bars show random regions as background. The y axis represents the total number of marks present in each category. Dashed lines in (F) and (H) indicate promoter and 5′ UTR regions. (I) Genes enriched in H3K4me3 or H3K27me3 marks are linked to distinct functional GO terms by ChIP-seq analysis. Bar graphs represent GO terms containing genes that have enrichment of the H3K4me3 mark (red scale) or the H3K27me3 mark (green scale) at their promoter regions in mIDH1-NS. The GO terms’ significance was determined by FDR (<0.05), and enrichment is expressed as odds ratio. (J and K) H3K4me3 occupancy in specific genomic regions of DNA repair regulatory genes Brat1 (J) and Rad51ap2 (K). The y axis of each profile represents the estimated number of immunoprecipitated fragments at each position normalized to the total number of reads in a given dataset. Reference sequence (RefSeq) gene annotations are shown. Differential peaks (FDR, <0.05) in mIDH1-NS are represented in red compared to wt-IDH1–NS in blue (n = 3 biological replicates per group).

The impact of H3 hypermethylation on epigenetic reprogramming was assessed by ChIP-seq, and the differential enrichment of H3K4me3 and H3K27me3 was evaluated (Fig. 2, B to H). The heat maps show differential peaks of histone marks centered at the peak midpoint in mIDH1-NS (cutoff, P < 1 × 10−5; Fig. 2, B and C, and fig. S7G). The average genomic distributions of H3K4me3 and H3K27me3 peaks in mIDH1-NS were around the transcription start sites (TSSs) (fig. S7H). Identified peaks were annotated to genomic features (Fig. 2D). We observed increased frequency of differential peaks for H3 marks in mIDH1-NS at promoters, 5′ untranslated region (UTRs), and around the first exon (Fig. 2, E to H). Because these regions are generally linked to transcriptional regulatory elements, our findings indicate that IDH1R132H could participate in epigenetic reprogramming. We identified enriched GO terms linked to differential H3K4me3 and H3K27me3 peaks (Fig. 2I). Gene promoters enriched for H3K4me3 are generally associated with transcriptional activation. Differential GO terms enriched in our model included DDR, cell cycle control, and regulation of cell development (Fig. 2I). Usually, gene promoters enriched in H3K27me3 are associated with transcription repression, and differential GO terms included cell differentiation. This would imply that IDH1R132H prevents differentiation, in agreement with the RNA-seq and immunohistochemistry (IHC) results (Fig. 1, D to G, and fig. S4). Differential peaks for H3K4me3 around the promoter regions of breast cancer type 1 (BRCA1)–associated ATM activator 1 (Brat1) and RAD51-associated protein 2 (Rad51ap2), which activate the HR repair pathway, were identified in mIDH1-NS, suggesting up-regulation of DDR mechanisms (Fig. 2, J and K).

IDH1R132H up-regulates expression of the ATM signaling pathway

RNA-seq analysis identified 1973 DE genes, including up-regulated genes related to “DNA damage stimulus responses” and “DNA repair” (fig. S8). GSEA indicated that DNA repair mechanisms were enriched in mIDH1-NS (FDR, <0.05; Fig. 3, A and B). Atm (Figs. 1D and 3B and fig. S8) and Brca1 (Fig. 1D and figs. S8 and S9) are key players in DNA repair and were shown to be DE in mIDH1-NS. This observation was mirrored in our enrichment maps showing biological functions involved in genomic stability: chromosome organization, DDR, DNA recombination, and cell cycle checkpoints (Fig. 3B and fig. S9). Fancd2, Rad50, and Rad51, which are involved in DNA repair and DDR, were up-regulated in mIDH1-NS (Fig. 1E and fig. S8).

Fig. 3 IDH1R132H up-regulates ATM signaling in the context of ATRX and TP53 KD.

(A) GSEA of transcriptional changes comparing mIDH1-NS versus wt-IDH1–NS. Positive normalized enrichment scores (red scale; FDR, <0.05) show GO terms linked to DDR and DNA repair pathways that are enriched in mIDH1-NS. (B) Pathway enrichment map of DE genes in mIDH1-NS versus wt-IDH1–NS. Clusters of red nodes illustrate differential enrichment (up-regulation) in mIDH1-NS (P < 0.05; overlap cutoff, >0.5), and they were extracted from the GSEA results comparing mIDH1-NS versus wt-IDH1–NS (fig. S3). The yellow highlighted circles indicate nodes containing Atm in up-regulated GO terms. (C) Expression of ATM, BRCA1, FANCD2, RAD50, and RAD51 in mouse NS. WB analysis using wt-IDH1–NS and mIDH1-NS. β-Actin, loading control. Bar graph represents semi-quantification of WB assay (n = 3). *P < 0.05; **P < 0.01; one-way ANOVA. Data shown as means ± SEM (n = 3 technical replicates). (D and E) WB for ATM and RAD51 performed on wt-IDH1 and mIDH1 SJGBM2 (D) and MGG8 (E) human glioma cells. Vinculin, loading control. Bar graph represents semi-quantification of WB assay (n = 3). *P < 0.05; **P < 0.01; one-way ANOVA. Bars represent means ± SEM (n = 3 technical replicates). (F) mRNA expression of DNA repair genes Brca1, Atm, Fancd2, and Rad51 in wt-IDH1–NS and mIDH1-NS. Reverse transcription–qPCR data are expressed relative to Hprt gene. *P < 0.05; **P < 0.01; ***P < 0.001; unpaired t test. Data are shown as means ± SEM (n = 3 technical replicates). (G and H) ChIP-qPCR for Atm (G) and Brca1 (H) was performed on isolated chromatin from wt-IDH1–NS and mIDH1-NS, immunoprecipitated with H3K4me3 and H3K27me3 antibodies. Diagram of Atm and Brca1 genomic regions indicates the qPCR primer positions (1, 2, and 3). Bar graphs show histone mark enrichment in the indicated genomic regions. Data are expressed as percentage of input and shown as means ± SEM. ****P < 0.0001; two-way ANOVA (n = 3 technical replicates). IgG, immunoglobulin G; Gapdh, glyceraldehyde-3-phosphate dehydrogenase.

WB analysis of NS demonstrated that expressions of ATM, BRCA1, FANCD2 [FA (Fanconi anemia) Complementation Group D2], RAD50, and RAD51 were increased in mIDH1-NS (Fig. 3C). In human glioma cells, IDH1R132H also increased expression of ATM and RAD51 (Fig. 3, D and E). IHC confirmed that RAD51, BRCA1, and ATM expression was enhanced in mIDH1 tumors, whereas p-DNA-PKcs (phosphorylated DNA-dependent protein kinase catalytic subunit) protein expression, involved in nonhomologous end-joining (NHEJ) repair, was not altered, and XRCC4 was decreased (fig. S10A). We validated these results by quantitative polymerase chain reaction (qPCR), observing increases in mRNA expression for Atm (2.8-fold; P < 0.05), Brca1 (7.6-fold; P < 0.01), Fancd2 (3.1-fold; P < 0.01), and Rad51 (5.3-fold; P < 0.01) in mIDH1-NS (Fig. 3F). Moreover, using ChIP-qPCR, we found that H3K4me3, but not H3K27me3, was significantly enriched in mIDH1-NS versus wt-IDH1–NS for both Atm (1.8-fold; P < 0.0001) (Fig. 3G) and Brca1 (2.2-fold; P < 0.0001) (Fig. 3H) genes around TSSs.

Gene expression analysis suggested that IDH1R132H could enhance HR repair and DDR (Fig. 4A). Thus, we performed a functional DNA repair assay (Fig. 4B) in human glioma cells (Fig. 4C) and NS expressing wt-IDH1 or mIDH1 (Fig. 4D). Human and mouse mIDH1 glioma cells exhibited enhanced HR repair efficiency [SJGBM2 (3.4-fold; P < 0.001), MGG8 (1.8-fold; P < 0.01), and mouse NS (1.6-fold; P < 0.01)] (Fig. 4, C and D). Treatment of wt-IDH1–NS with 2.5 mM (2R)-octyl-α-hydroxyglutarate (O-2HG), a cell-permeable analog of 2HG, enhanced HR repair efficiency (Fig. 4E).

Fig. 4 IDH1R132H enhances DNA repair efficiency and confers in vitro radioresistance.

(A) Diagram of HR repair pathway. (B) HR repair reporter assay. Diagram shows the HR reporter plasmid and the mechanism to measure HR repair efficiency by reconstitution of green fluorescent protein (GFP) expression. (C to E) Bar graphs show HR repair efficiency in wt-IDH1 and mIDH1 human glioma cells (C), wt-IDH1–NS and mIDH1-NS (D), and wt-IDH1–NS treated with O-2HG (E). GFP expression was normalized to blue fluorescent protein (BFP) expression. **P < 0.01; ***P < 0.001; unpaired t test (n = 3 technical replicates; data are shown as means ± SEM). (F) Quantification of γH2AX foci in wt-IDH1–NS and mIDH1-NS from 0 to 48 hours after IR (2 Gy; fig. S9). Bar graph represents the average number of foci per nuclei ± SEM. ***P < 0.001; ****P < 0.0001; one-way ANOVA (n = 3 technical replicates). (G) Quantification of 53BP1 foci in wt-IDH1–NS and mIDH1-NS from 0 to 48 hours after IR (2 Gy). Bar graph represents the average number of foci per nuclei ± SEM. ***P < 0.001; ****P < 0.0001; one-way ANOVA (n = 3 technical replicates). (H) WB analysis shows γH2AX, pCHK2, pRPA32, pATM, and their respective nonphosphorylated proteins from 0 to 48 hours after IR (2 Gy). Vinculin: loading control. (I) WB analysis shows γH2AX, pRPA32, and pATM from 0 to 48 hours after IR (20 Gy). Vinculin, loading control. (J) Diagram of Bru-seq assay to identify nascent RNAs labeled with Bru. (K and L) Bru-seq traces show differential transcriptional rates (<1.5-fold; P < 0.05) of DNA repair genes Atm (K) and Rad50 (L) in mIDH1-NS (orange) compared to wt-IDH1–NS (blue). Arrows indicate sequence strand reading direction. Atm and Rad50 are shown at the top in red. The positive y axis represents the positive strand signal of transcription moving from left to right, and the negative y axis represents the negative strand signal of transcription moving from right to left. The vertical red mark indicates the position of Atm and Rad50 within their respective chromosomes. The gene maps were generated from RefSeq. Genes and chromosome locations are indicated on the maps. Data are expressed in reads per kilobase per million mapped reads (RPKM). (M to O) Impact of mIDH1 on radiosensitivity in NS: NRAS/shP53/shATRX (M), PDGFB/shP53/shATRX/Ink4a/Arf−/− (N), and human glioma cells SJGBM2 (O). Cell viability assay shows the effect of AGI-5198 ± IR on cell proliferation in wt-IDH1 and mIDH1 mouse and human glioma cells. Results are expressed in relative luminescence units (RLU). **P < 0.01; ****P < 0.0001; two-way ANOVA (n = 3 technical replicates).

We next quantified the kinetics of γH2AX and 53BP1 foci formation in response to ionizing radiation (IR) (Fig. 4, F and G). The formation of γH2AX foci was increased ~2-fold versus basal quantities (P < 0.0001) 0.5 hours after IR in both wt-IDH1 and mIDH1-NS, indicating DNA damage and DDR activation (Fig. 4F). At 4 hours after IR, the number of foci in wt-IDH1–NS continued to increase (~2.5-fold; P < 0.0001), whereas in mIDH1-NS, the number of foci significantly decreased (~30%; P < 0.0001). At 48 hours, the number of foci in mIDH1 returned to baseline (Fig. 4F and fig. S10B). Similarly, the average number of 53BP1 foci per cell increased at 0.5 hours after IR in both wt-IDH1–NS and mIDH1-NS (P < 0.0001; Fig. 4G). However, mIDH1-NS reached basal foci number before wt-IDH1–NS, indicating faster double-strand break (DSB) repair. In addition, we performed a neutral comet assay to assess genome integrity and DSB repair kinetics after IR (fig. S10C). IR immediately generated DNA damage characterized by increased nuclear tail lengths, proportional to the number of DSBs at neutral pH. Scores were proportionately higher (longer tails) for wt-IDH1, indicating greater DNA damage at earlier time points (P < 0.05; fig. S10C).

We next studied the phosphorylation status of γH2AX, pCHK2, pRPA32, and pATM after IR. WB data showed a peak of γH2AX 0.5 hours after IR in both wt-IDH1–NS and mIDH1-NS (Fig. 4H). However, the γH2AX signal decreased in mIDH1-NS at 4 hours after IR compared to wt-IDH1–NS, indicating that mIDH1-NS repaired their DNA more efficiently (Fig. 4H). Similarly, pCHK2, pRPA32, and pATM signals decreased after 4 hours. In wt-IDH1, these signals remained positive 48 hours after IR (Fig. 4H). We also observed increased expression of total ATM in mIDH1-NS 4 hours after IR. Human glioma cells expressing IDH1R132H displayed a similar pattern (Fig. 4I and fig. S10, D and E). Our data show that DNA repair efficiency is enhanced in mIDH1 tumor cells, in agreement with our RNA-seq and CHIP-seq analyses (Figs. 1E and 2I). This was also assessed by Bru-seq analysis (Fig. 4J). mIDH1-NS display higher transcription rate for Atm and Rad50 (>1.5-fold) compared to wt-IDH1–NS (Fig. 4, K and L). Cell viability, evaluated after IR, was higher in mIDH1-NS versus wt-IDH1–NS (~2-fold; P < 0.0001) (Fig. 4M). In addition, mIDH1-NS showed decreased NHEJ repair activity (fig. S10F), an error-prone DNA repair mechanism.

RAS pathway activation can confer radioresistance (19); thus, we validated our results using NS generated from a mIDH1 glioma model independent of RAS-activating mutations (20). Brain tumors were induced with replication-competent avian leukemia virus splice acceptor (RCAS) platelet derived growth factor β (PDGFB), mIDH1 or wt-IDH1, and shP53 in mixed background NTva, Ink4a/Arf−/− mice (20). We engineered the NS to encode shATRX to generate glioma cells with the following molecular alterations: PDGFB/shP53/shATRX/Ink4a/Arf−/−/mIDH1 or PDGFB/shP53/shATRX/Ink4a/Arf−/−/wt-IDH1. Using this model, we confirmed that mIDH1 confers radioresistance (~2-fold; P < 0.0001) (Fig. 4N). Likewise, human glioma cells harboring IDH1R132H displayed higher viability in response to IR (2.3-fold; P < 0.0001) (Fig. 4O). These results were further validated in human glioma cells with endogenous expression of IDH1R132H in the context of ATRX and TP53 inactivation (SF10602 and LC1035), which showed increased expression of RAD51 and ATM (fig. S11, A and B) and also displayed radioresistance (fig. S11, C and D). After treatment with AGI-5198, both mIDH1-NS and mIDH1 human glioma cells (Fig. 4, M to O, and fig. S11, E to G) became radiosensitive. In vitro radiosensitivity was also evaluated by clonogenic survival assay, with the survival dose-response curve fitted to a linear regression (fig. S12). This experiment confirmed that mouse and human mIDH1 cells are less sensitive to radiation. The enhanced DDR in mIDH1 cells correlated with faster digestion of chromatin, indicating that mIDH1 cells have less condensed chromatin at a global level (fig. S13), which could facilitate recruitment of the DDR response machinery to DNA damage sites. Collectively, these results indicate that IDH1R132H enhances DDR, imparting radioresistance independently of the presence of RAS-activating mutations.

IDH1R132H confers radioresistance in intracranial glioma models

To investigate the in vivo effects of mIDH1 in response to IR, we designed a preclinical trial using an intracranial glioma model, generated by implanting wt-IDH1–NS and mIDH1-NS into the brain of adult mice (Fig. 5A). At 7 days post-implantation (DPI) with wt-IDH1–NS or mIDH1-NS, animals were treated with IR at the indicated doses. Untreated animals harboring wt-IDH1 tumors exhibited a MS = 21 days, which was increased after 20 grays (Gy) IR (MS = 51 days; P = 0.0005) (Fig. 5B). In contrast, animals harboring mIDH1 tumors exhibited MS = 33 days, which did not increase in response to IR (Fig. 5C). These results demonstrate that IDH1R132H confers radioresistance in vivo. We then compared the genome-wide gene expression profiles using RNA-seq after IR (Fig. 5, D to I). We examined the following animal groups: (i) IR wt-IDH1 tumors (wt-IDH1-R) versus nontreated (NT) wt-IDH1 tumors (wt-IDH1-NT) (Fig. 5D) and (ii) IR mIDH1 tumors (mIDH1-R) versus NT mIDH1 tumors (mIDH1-NT) (Fig. 5E). We found that the number of DE genes between mIDH1-R tumors and mIDH1-NT tumors increased at 20 Gy compared to 10 Gy (Fig. 5F). Wt-IDH1 tumors had fewer DE genes compared to mIDH1 tumors at both 10 and 20 Gy (Fig. 5F). Moreover, wt-IDH1-R tumors did not exhibit an increase of DE genes between 10 and 20 Gy (Fig. 5F). This suggests that in vivo resistance to radiation-induced DNA damage in mIDH1 glioma involves differential gene expression. Functionally, the genes up-regulated in mIDH1 gliomas in response to IR are linked to regulation of cell proliferation, cell migration, and cell homeostasis (Fig. 5, G and H). In addition, several genes involved in DNA repair were up-regulated in mIDH1-R (Fig. 5I and fig. S14), indicating that inducible DNA repair mechanisms were associated with in vivo radioresistance in the mIDH1 glioma model. We analyzed the in vivo expression of Ki-67 (proliferation), CC3 (apoptosis), and γH2AX (DNA damage) at 14 and 21 DPI either in animals treated with 10 or 20 Gy or in untreated animals (Fig. 5, A and J to N). In wt-IDH1-NT mice, we observed that Ki-67 (Fig. 5, J and L) significantly decreased from 14 to 21 DPI (>105-fold; P < 0.0001). Presumably, the wt-IDH1 tumors had reached their maximum size (Fig. 5O), and tumor cells were no longer proliferating because the MS for untreated animals harboring wt-IDH1 tumors is 21 days. Conversely, after 20 Gy, Ki-67 expression was enhanced (>105-fold; P < 0.0001), correlating with proliferating tumor cells. In mIDH1 mice, we observed decreased Ki-67 at 21 DPI compared to 14 DPI but no differences between the IR and NT groups (Fig. 5, K and L), consistent with the lack of effect of IR on MS (Fig. 5C) and tumor size (Fig. 5P). CC3 was significantly higher in wt-IDH1-R at 14 and 21 DPI compared to the NT group (>105-fold; P < 0.0001) (Fig. 5, J and M), which correlated with a reduction in tumor size (Fig. 5O). In mIDH1 tumors, CC3 expression was low in all experimental groups (Fig. 5, K and M), indicating no IR-mediated tumor cell death. In addition, γH2AX increased in both wt-IDH1 and mIDH1 tumors treated with 10 and 20 Gy, indicative of DNA damage foci (Fig. 5N). However, in wt-IDH1 tumors, γH2AX increased at 21 DPI versus 14 DPI (7.5-fold; P < 0.01), whereas in mIDH1, it decreased at 21 DPI versus 14 DPI (4.5-fold; P < 0.05), implying better DNA repair activity (Fig. 5N). Together, these results suggest that IDH1R132H induces radioresistance in vivo by altering gene expression, enhancing DDR and DNA repair mechanisms.

Fig. 5 IDH1R132H confers in vivo radioresistance.

(A) Preclinical trial design for testing the effect of mIDH1 on radio response in an orthotopic mIDH1 glioma model. mIDH1-NS and wt-IDH1–NS were implanted into adult mice (day 0). At seven DPI, animals were split into six groups: (i) 2 Gy/day for 5 days (total = 10 Gy), euthanized at 14 DPI (n = 7); (ii) NT group, euthanized at 14 DPI (n = 5); (iii) 2 Gy/day for 10 days (total = 20 Gy), euthanized at 21 DPI (n = 7); (iv) NT, euthanized at 21 DPI (n = 5); (v) 2 Gy/day for 10 days (total = 20 Gy), euthanized at moribund stage (MS = 51 DPI for wt-IDH1 and 38 DPI for mIDH1 tumor–bearing mice; n = 7); and (vi) NT, euthanized at moribund stage (MS = 21 DPI for wt-IDH1 and 33 DPI for mIDH1 tumor–bearing mice; n = 5). Tumors were processed for IHC and RNA-seq analyses. (B and C) Kaplan-Meier survival curves of (B) wt-IDH1 and (C) mIDH1 tumor–bearing mice either treated with IR (20 Gy; n = 7) or NT (n = 5). Statistical significance was determined by Mantel-Cox test. (D) Volcano plot showing the comparison of gene expression in wt-IDH1 tumors from mice treated with IR (20 Gy) and processed at moribund stage (wt-IDH1-R) versus untreated wt-IDH1 tumors (wt-IDH1-NT). The −log10 q values were plotted against the log2 fold change. Up-regulated genes (n = 55; ≥1.5-fold; FDR-corrected P <0.05) are depicted as red dots, and genes that were down-regulated (n = 149; ≥1.5-fold; FDR-corrected P < 0.05) are depicted as green dots. FDR-corrected P values = q values; two-sided moderated Student’s t test. (E) Volcano plot showing the comparison of gene expression in mIDH1 tumors from mice treated with IR (20 Gy) and processed at moribund stage (mIDH1-R) versus control mIDH1 tumors that were not treated (mIDH1-NT). The −log10 q values were plotted against the log2 fold change. Up-regulated genes (n = 1295; ≥1.5-fold; FDR-corrected P <0.05) are depicted as red dots, and genes that were down-regulated (n = 184; ≥1.5-fold; FDR-corrected P <0.05) are depicted as green dots. FDR-corrected P values = q values; two-sided moderated Student’s t test. (F) Bar graph showing the total number of DE genes (≥1.5-fold; q < 0.05) in wt-IDH1 (wt-IDH1-R versus wt-IDH1-NT) and mIDH1 (mIDH1-R versus mIDH1-NT) gliomas after 10 or 20 Gy. (G) Functional enrichment of DE genes in mIDH1-R group. (H) Pathway enrichment map from GSEA of mIDH1-R tumors versus mIDH1-NT tumors. Red nodes illustrate differential enrichment (up-regulation) in mIDH1-R tumors (P < 0.05; overlap cutoff, >0.5). (I) Heat map showing gene expression pattern for DNA repair genes in mIDH1-NT and mIDH1-R tumors treated with 20 Gy (n = 3 biological replicates). Differentially up-regulated genes are depicted in red, whereas down-regulated genes are depicted in green (FDR, ≤0.05; ≥1.5-fold). (J and K) Immunofluorescence staining of Ki-67, CC3, and γH2AX in wt-IDH1 (J) and mIDH1 (K) mice at 14 and 21 DPI ± IR treatment at the indicated doses. (L to N) Quantification of immunofluorescence staining. Bar graphs represent total numbers of cells positive for Ki-67 (L), CC3 (M), and γH2AX (N). *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001; unpaired t test. Data are shown as means ± SEM (n = 3 biological replicates). (O and P) Tumor progression was evaluated by tumor size quantification of wt-IDH1 (O) and mIDH1 (P) tumor sections at 14 and 21 DPI with or without IR treatment at the indicated doses. Tumor size is expressed as percentage relative to untreated tumors at 14 or 21 DPI (100%).

Inhibition of DDR pathways restores radiosensitivity in mIDH1 glioma

The increased DDR and radioresistance observed in mIDH1 glioma suggest that pharmacological DDR inhibition could improve the response to IR. In vitro cell viability assays showed decreased sensitivity of mIDH1-NS to IR for both mouse glioma models (Fig. 6, A and B). We observed that temozolomide (TMZ), which is standard of care for patients with glioma, confers radiosensitivity in mIDH1-NS and mIDH1 human glioma cells (fig. S15, A to C). Similarly, IR combined with specific inhibitors for ATM (KU60019) (Fig. 6, A and B) or checkpoint kinases 1 and 2 (CHK1/2; AZD7762) (Fig. 6, C and D) decreased cell viability (P < 0.0001). Comparable results were observed in human glioma cells (fig. S15, D to I). To assess in vivo DDR inhibition in response to IR, we used the intracranial mIDH1 model described above. Animals were treated with IR combined with KU60019 (Fig. 6E). Radiation or ATM inhibition alone did not modify MS compared to NT animals (Fig. 6F). However, KU60019 combined with IR improved MS of mIDH1 mice (45 days) versus controls (MS = 30 days; P < 0.01) (Fig. 6F), consistent with decreased tumor size in mIDH1 animals treated with 20 Gy and KU60019 (Fig. 6G). To study the effect of cell cycle progression in response to IR in mIDH1 glioma, we combined IR with CHK1/2 inhibition in vivo (Fig. 6H). AZD7762 combined with IR increased MS in mIDH1 glioma–bearing mice (Fig. 6I). Tumor size was also significantly decreased in the group treated with combined IR and AZD7762 at 14 DPI (5-fold; P < 0.0001) and 21 DPI (11-fold; P < 0.0001) (Fig. 6J). In wt-IDH1 tumors, ATM inhibition with IR did not improve MS compared to IR alone (fig. S15J). However, CHK1/2 inhibition with IR increased MS in wt-IDH1 glioma–bearing mice (fig. S15K). We also assessed CC3 expression in mIDH1 tumors after treatment with IR combined with KU60019 or AZD7762 at 14 DPI (Fig. 6, K and L). CC3 expression increased in mIDH1 tumors treated with IR combined with KU60019 or AZD7762, suggesting that DDR inhibition in combination with IR induced apoptosis. In mIDH1 human glioma cells (SF10602 and LC1035), AZD7762 reverted radioresistance (Fig. 6, M and N). Consistent with our results, analysis of The Cancer Genome Atlas (TCGA) database indicated that patients with LGG harboring IDH1R123H with TP53 and ATRX inactivating mutations have higher expression of ATM and RAD50 mRNA than wt-IDH1 GBM (fig. S15, L and M) and higher ATM expression than patients with wt-IDH1-LGG (fig. S15N). In addition, the up-regulation of ATM in patients with glioma correlated with increased survival (fig. S15O).

Fig. 6 Inhibition of DDR reverts in vivo radioresistance in mIDH1 glioma.

(A and B) Inhibition of ATM pathway in mouse glioma cells expressing mIDH1. In vitro data showing cell proliferation of mouse NS NRAS/shP53/shATRX (A) and PDGFB/shP53/shATRX/Ink4a/Arf−/− (B) with or without mIDH1 in response to IR (6 Gy) with or without 1.5 μM of KU60019. ****P < 0.0001; two-way ANOVA. Data are shown as means ± SEM (n = 3 technical replicates). (C and D) Inhibition of CHK1/2 in mouse mIDH1-NS. In vitro data showing cell proliferation of mouse NS NRAS/shP53/shATRX (C) and PDGFB/shP53/shATRX/Ink4a/Arf−/− (D) with or without mIDH1 in response to IR (6 Gy) with or without 1.5 μM of AZD7762. ****P < 0.0001; two-way ANOVA. Data are shown as means ± SEM (n = 3 technical replicates). (E) Preclinical trial design for testing the impact of the ATM inhibitor (KU60019) on the response to IR in an orthotopic glioma model. Seven days after implantation of NS, animals were separated into eight groups: (i) untreated (NT), euthanized at 14 DPI; (ii) IR 2 Gy/day for 5 days (total = 10 Gy), euthanized at 14 DPI; (iii) KU60019 (continuous infusion for 5 days; 36 mg/kg per day), euthanized at 14 DPI; (iv) KU60019 (continuous infusion for 5 days; 36 mg/kg per day) and 2 Gy/day for 5 days (total = 10 Gy), euthanized at 14 DPI; (v) NT, euthanized at moribund stage (between 30 and 50 DPI); (vi) 2 Gy/day for 10 days (total = 20 Gy), euthanized at moribund stage; (vii) KU60019 (continuous infusion for 10 days; 36 mg/kg per day), euthanized at moribund stage; and (viii) KU60019 (continuous infusion for 10 days; 36 mg/kg per day) and 2 Gy/day for 10 days (total = 20 Gy), euthanized at moribund stage. (F) Kaplan-Meier survival curve of mIDH1 tumor–bearing mice treated with or without 20 Gy (n = 5) in the presence or absence of KU60019 (n = 6). **P < 0.01; Mantel-Cox test. (G) Tumor progression evaluated by tumor size of mIDH1 glioma sections at 14 and 21 DPI ± IR at the indicated doses in the presence or absence of KU60019. Tumor size was expressed as percentage relative to untreated tumors at 14 DPI (100%) ± SEM. **P < 0.01; one-way ANOVA. (H) Trial design for testing the impact of CHK1/2 signaling inhibitor (AZD7762) on the response to IR in an orthotopic glioma model. Seven days after implantation of NS, the animals were separated into eight groups as described in (E) but treated with AZD7762 (15 mg/kg per day). (I) Kaplan-Meier survival curve of mIDH1 tumor–bearing mice ± 20 Gy (n = 5), with or without AZD7762 (15 mg/kg per day; n = 6). MS of mice bearing mIDH1 tumors significantly increased after IR combined with AZD7762. *P < 0.01; Mantel-Cox test. (J) Tumor size evaluated by quantification of mIDH1 brain tumor sections at 14 and 21 DPI ± IR at the indicated doses, in the presence or absence of AZD7762. Tumor size is expressed as percentage relative to untreated tumors at 14 DPI (100%) ± SEM. ***P < 0.001; ****P < 0.0001 one-way ANOVA. (K) Immunofluorescence staining of CC3 expression at 14 DPI ± IR treatment at the indicated doses, in the presence or absence of KU60019 or AZD7762. (L) Bar graphs represent total numbers of CC3+ cells in (K). ****P < 0.0001; unpaired t test. Data are shown as means ± SEM. (M and N) Impact of CHK1/2 inhibition on radioresistance in human glioma cells with endogenous expression of mIDH1: SF10602 (M) and LC1035 (N). Cell viability assay shows the effect of AZD7762. The results are expressed in RLU. *P < 0.05; **P < 0.01; ****P < 0.0001; two-way ANOVA.

DISCUSSION

Patients harboring IDH1R132H glioma exhibit longer MS (~6.6 years from diagnosis) compared with patients whose tumors express wt-IDH1 (~1.6 years from diagnosis) (2, 3, 21). In line with this, our mIDH1 mouse model exhibits increased MS greater than twofold compared with wt-IDH1 tumors. It was reported (5) that the glioma subgroup harboring IDH1R132H, ATRX, and TP53 loss also exhibits lengthening of telomeres. In our mouse glioma model, telomere elongation is mediated by ALT (4, 22). Genomic stability in our mIDH1 glioma model is mediated via increased DDR due to epigenetic reprogramming of the tumor cells’ transcriptome (fig. S16). DDR disruption is one of the hallmarks of gliomas and other cancers (23). ATM kinase senses DSB lesions on the DNA, activating responses that maintain genome integrity (24). Our ChIP-seq data revealed enrichment of H3K4me3 at promoter regions of genes involved in DDR and cell cycle progression. ChIP-qPCR showed enrichment of the H3K4me3 mark around Atm TSSs, which would increase Atm expression. This was confirmed by Bru-seq and RNA-seq studies. ATM was also up-regulated in patients with LGG harboring IDH1R132H with ATRX and TP53 gene inactivation; this correlated with increased survival of these patients. We discovered that IDH1R132H induced transcriptional activation of Atm, which resulted in efficient DNA repair activity via HR repair (25, 26). Because the chromatin was less condensed in mIDH1 glioma cells, this could enable the recruitment of the DDR machinery to sites of DNA damage.

Mutations in IDH1/2 are also detected in 15% of patients with acute myeloid leukemia (AML), which correlates with unfavorable prognosis (27). ATM expression is down-regulated (through H3K9me3), and DDR functions and genomic stability are also reduced in this setting (28). mIDH1 AML cells are more sensitive to chemotherapy and are highly malignant. Thus, the production of 2HG has opposite effects in these two different cancers. This highlights the critical influence of the genetic context in which IDH1R132H acts. In the glioma subtype that we studied, IDH1R132H is expressed in the context of TP53 and ATRX mutations, differing from AML, where ATRX gene inactivation is not present (27, 29).

We hypothesized that the increase in DDR elicited by mIDH1 could induce radioresistance because mIDH1 glioma cells are able to more efficiently repair DNA damage inflicted by IR. It appears that IDH1R132H induces genomic stability, which on one hand slows tumor growth and, on the other, increases DNA repair capacity, reducing the efficacy of radiotherapy. Previous studies using colon cancer cells, HeLa cells, and cells derived from high-grade gliomas with ectopic expression of mIDH1 suggest that IDH1R132H renders these cells more sensitive to radiation (3032). None of these cells, however, originated from patient-derived IDH1R132H glioma, harboring concomitant mutations in ATRX and TP53, and no experiments were done orthotopically. Note that Sulkowski et al. (33) studied primary patient-derived glioma cultures with IDH1 mutations and demonstrated clear DNA repair deficiency and sensitivity to poly(adenosine 5′-diphosphate–ribose) polymerase inhibition. Their results suggest that at least some subsets of human mIDH1 gliomas may be deficient in DNA repair and, consequently, radiosensitive. These apparently opposing results reinforce the notion that the effects of IDH1R132H can vary according to tumor type/subtype and should be evaluated in an appropriate cellular and genetic context. Our results indicating that IDH1R132H decreases radiosensitivity and enhances DDR in glioma were validated in cells derived from patients with glioma with endogenous expression of IDH1R132H in the context of TP53 and ATRX inactivation and in a second mouse glioma model lacking RAS-activating mutation (20). In agreement with our results, a recent study using gliomaspheres demonstrated that mIDH1 cultures are less sensitive to IR than wt-IDH1 cultures; however, this work did not distinguish between 1p/19q codeleted and non-codeleted mIDH1 glioma subtypes (34).

Thus, we postulated that inhibiting DDR would restore glioma radiosensitivity in the mIDH1 glioma subtype under investigation. When we blocked DDR, the tumors’ sensitivity to IR therapy was restored (fig. S16). In patients, the effect of IDH1R132H on IR response remains controversial. Evidence in favor of IDH1R132H increasing or decreasing tumors’ radiosensitivity has been published (21, 3537). Patients with glioma expressing IDH1R132H do live longer, but whether this is due to IDH1R132H tumors growing slower or whether they are more radiosensitive has not yet been conclusively demonstrated. To conclusively demonstrate sensitivity to radiation in patients with mIDH1 glioma, a control group not treated with radiation would be needed. Our in vivo results indicate that mIDH1 glioma–bearing mice do not exhibit a therapeutic response to IR. Our data also demonstrate that the effects mediated by IDH1R132H on DDR are dependent on the genetic context. This is in agreement with the only study available in the literature, which included 300 patients with LGG treated with or without radiotherapy (37). Similarly, survival of WHO II patients with glioma expressing IDH1R132H treated with TMZ was not further improved by radiotherapy (35). In addition, the combination of radiation therapy followed by vincristine, procarbazine, and lomustine in WHO II glioma prolonged overall survival compared with patients receiving IR alone (36), suggesting that tumors expressing IDH1R132H remain sensitive to chemotherapy but not radiotherapy.

Although the results we reported have therapeutic implications for patients harboring mIDH1 LGGs, there are nevertheless, some limitations to our work, which one has to take into consideration. First, our genetically engineered mIDH1 mouse glioma model, in addition to IDH1R132H, ATRX, and TP53 inactivation, also harbors activated NRAS G12V. Although it has been previously shown that the RTK/RAS/PI3K pathway is activated in gliomas (38, 39), activated NRAS mutations have not been reported in human gliomas, on the basis of TCGA and other human databases (38, 39). Second, although we did demonstrate increases in DDR and radioresistance ex vivo, in primary patient-derived mIDH1 LGG cell cultures harboring ATRX and TP53 loss, because these cells failed to form tumors in immunodeficient mice, we were unable to test their sensitivity to radiation in vivo. Nevertheless, using a second mouse glioma model of the desired genotype, which does not harbor RAS-activating mutations, and several patient-derived mIDH1 LGG cell cultures, we present consistent data, which point to increased HR resulting in radioresistance in this mIDH1 LGG subtype. In addition, DDR inhibitors, both in vitro and in vivo, conferred increased sensitivity to radiation against this mIDH1 glioma subtype, lending support to eventual clinical testing of this therapeutic combination.

In conclusion, we discovered the mechanism by which IDH1R132H, in the context of TP53 and ATRX inactivation, elicits epigenetic reprogramming of the ATM signaling pathway, which in turn increases DDR and genomic stability (fig. S16). Our data suggest combining radiation with DDR inhibition to increase therapeutic efficacy in patients with mIDH1 LGG.

MATERIALS AND METHODS

Study design

To study the impact of IDH1R123H activity in the context of TP53 and ATRX KD, we generated a genetically engineered animal model injecting SB plasmids encoding NRAS G12V and shp53, with or without shATRX and with or without IDH1R132H into the lateral ventricle of neonatal mice. Sample size and any data inclusion/exclusion were defined individually for each experiment. In addition, we used an animal model generated by intracranial implantation of glioma NS (wt-IDH1 and mIDH1) derived from our genetically engineered animal model to test therapeutic responses. We also used an alternative model (PDGFB/shP53/shATRX/Ink4a/Arf−/− wtIDH1-NS or mIDH1-NS) that does not encode RAS-activating mutations. In addition, we used human glioma cells derived from patients harboring IDH1R132H in the context of TP53 and ATRX inactivating mutations to confirm the results obtained from our animal models. The numbers of replicates are reported in the figure legends. Our studies were not randomized. We performed blinding for quantitative IHC scoring. All RNA-seq and ChIP-seq data were deposited in public databases as is indicated in the respective sections. Materials and Methods are detailed in the Supplementary Materials.

Statistical analysis

All quantitative data are presented as the means ± SEM from at least three independent samples. ANOVA and two-sample t tests were used to compare continuous outcomes between groups. Survival curves were analyzed using the Kaplan-Meier method and compared using Mantel-Cox tests; the effect size is expressed as MS. Differences were considered significant if P < 0.05. All analyses were conducted using GraphPad Prism software (version 6.01), SAS (version 9.4, SAS Institute), or R (version 3.1.3). The statistical tests used are indicated in each figure legend.

SUPPLEMENTARY MATERIALS

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Materials and Methods

Fig. S1. Generation of mIDH1 mouse glioma model using the SB transposon system.

Fig. S2. Characterization of tumor NS generated from wt-IDH1 and mIDH1 mouse brain tumors and human glioma.

Fig. S3. Functional enrichment of DE genes in mIDH1-NS versus wt-IDH1–NS.

Fig. S4. Tumor cell differentiation phenotype in mIDH1 glioma.

Fig. S5. TICs in wt-IDH1 and mIDH1 glioma cells.

Fig. S6. In vivo cell cycle analysis on wt-IDH1 and mIDH1 tumors.

Fig. S7. Analysis of histone modifications in mIDH1-NS.

Fig. S8. DE genes in mIDH1-NS determined by RNA-seq analysis.

Fig. S9. Pathway enrichment map of differential gene expression in mIDH1 versus wt-IDH1 mouse NS.

Fig. S10. Analysis of DNA repair efficiency and DDR activity in wt-IDH1 and mIDH1 glioma.

Fig. S11. In vitro radioresistance analysis in wt-IDH1 and mIDH1 glioma cells.

Fig. S12. Clonogenic assay performed on wt-IDH1 and mIDH1 mouse and human glioma cells.

Fig. S13. Chromatin compaction in wt-IDH1 and mIDH1 cells.

Fig. S14. Heat map showing gene expression for DNA repair pathways in mIDH1-NT and mIDH1-R (20 Gy).

Fig. S15. Impact of DDR activity on in vitro radio response in mIDH1 glioma cells.

Fig. S16. Diagram of epigenetic reprograming elicited by mIDH1 on DDR pathway up-regulation, survival, and response to radiotherapy in glioma.

Table S1. Antibodies.

Table S2. Oligonucleotides.

References (4051)

REFERENCES AND NOTES

Acknowledgments: We thank E. Holland for providing RCAS IDH1 R132H, RCAS IDH1 WT plasmids (Fred Hutchinson Cancer Research Center and the University of Washington, Seattle); J. Ohlfest (University of Minnesota, deceased) for providing the SB model plasmids; J. Costello (UCSF); and H.Wakimoto and D. Cahill (Harvard Medical School) for providing mIDH1 human glioma cells, SF10602 and MGG119, respectively. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH. Funding: This work was supported by NIH/NINDS grants R37-NS094804 and R01-NS105556 to M.G.C., NIH/NINDS grants R01-NS076991 and R01-NS096756 to P.R.L., NIH/NIBIB grant R01-EB022563, NIH/NCI grant U01CA224160, the Department of Neurosurgery and Leah’s Happy Hearts to M.G.C. and P.R.L., RNA Biomedicine grant F046166 to M.L. and M.G.C., NIH/NCI grant T32-CA009676 to S.C., NIH/NINDS grant 5K08NS099427-02 to C.K., NIH/NINDS grant 1F31NS103500 to F.M.M., Aflac Cancer and Blood Disorders Center to D.H., NIH/NINDS grant F31NS106887 to C.H., University of Michigan's Program in Chemical Biology Graduate Assistance in Areas of National Need (GAANN) to D.M.K., and 2017 AACR NextGen Grant for Transformative Cancer Research (17-20-01-LYSS) to C.A.L. Metabolomics studies were supported by NIH grant DK097153, the Charles Woodson Clinical Research Fund, and the UM Pediatric Brain Tumor Initiative to C.A.L; Dabbiere family and NIH grants 5T32CA151022-07 and 5R01CA169316-05 to L.J.; NIH grant K08 CA181475 to S.V.; and U.S. National Institute of Aging to V.G. Author contributions: F.J.N., P.R.L., and M.G.C. conducted and designed the experiments, analyzed the data, and wrote the manuscript. F.M.M., P.K., M.S.A., M.G.S., C.K., A.-A.C., N.K., M.S., R.P., S.C., M.Z.G., M.B.G.-F., S.H., and M.E., performed the experiments and data analysis. M.L. conducted Bru-seq analysis. T.Q. and M.A.S. performed ChIP-seq analysis. R.T. performed RNA-seq analysis. J.B.-C. and A.M. performed the ALT experiments. V.G. contributed with DNA repair reporter plasmids. L. Zhao performed the statistical analysis of the data. S.H.-J. and S.V. contributed to the IHC analysis of human samples. D.M.K., L. Zhang, and C.A.L. evaluated the 2HG concentration. C.J.H., J.L.R., and D.H. generated the glioma mouse models using the RCAS system. L.J. generated and characterized the SF10602 human glioma cells. M.E.F. contributed to the ChIP-seq experimental design and data analysis. P.R.L. and M.G.C. directed the research and generated the funding. All authors read and edited the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data associated with this study are in the paper and/or the Supplementary Materials. The RNA-seq datasets have been deposited in NCBI's Gene Expression Omnibus with identifiers GSE94902, GSE94974, and GSE94975. The ChiP-seq datasets have been deposited in NCBI's Gene Expression Omnibus with identifier GSE99806. The materials generated in this study can be requested from M.G.C.
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