Research ArticleCANCER TREATMENT

Targeting the kinase activities of ATR and ATM exhibits antitumoral activity in mouse models of MLL-rearranged AML

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Sci. Signal.  13 Sep 2016:
Vol. 9, Issue 445, pp. ra91
DOI: 10.1126/scisignal.aad8243

New hope for AML patients

A pair of papers provides new hope for patients with acute myeloid leukemia (AML) by showing that the DNA replication checkpoint pathway is a viable target for therapeutic intervention. By integrating survival data from 198 treated AML patients with gene expression data for genes encoding proteins involved in the regulation of DNA replication, David et al. identified the CHEK1 gene and its product, the DNA replication checkpoint kinase CHK1, as both a prognostic indicator of survival and a therapeutic target to overcome resistance to the current standard of chemotherapy. The patients had all received standard-of-care chemotherapy. Patients with high expression of CHEK1 in their AML cells had reduced survival, and AML patient cells with high CHK1 abundance were resistant to the toxic effects of the DNA replication inhibitor cytarabine. CHK1 is activated by the kinase ATR in response to DNA replication stress arising from DNA damage. The identification of CHEK1 expression as high in lymphomas and leukemias, including AML, prompted Morgado-Palacin et al. to investigate targeting ATR and ATM, the most upstream kinases in the DNA damage response, as possible AML therapies. AML cells with oncogenic rearrangements in MLL are particularly resistant to genotoxic therapies that form the backbone of AML treatment. Inhibiting ATR resulted in death of AMLMLL cells in culture and exhibited antitumoral activity in AMLMLL mouse models. Inhibiting ATM also prolonged survival of the allograft mouse model, indicating that targeting the DNA damage response pathways alone or in combination with other chemotherapeutic agents may be beneficial in patients with AML.

Abstract

Among the various subtypes of acute myeloid leukemia (AML), those with chromosomal rearrangements of the MLL oncogene (AML-MLL) have a poor prognosis. AML-MLL tumor cells are resistant to current genotoxic therapies because of an attenuated response by p53, a protein that induces cell cycle arrest and apoptosis in response to DNA damage. In addition to chemicals that damage DNA, efforts have focused on targeting DNA repair enzymes as a general chemotherapeutic approach to cancer treatment. Here, we found that inhibition of the kinase ATR, which is the primary sensor of DNA replication stress, induced chromosomal breakage and death of mouse AMLMLL cells (with an MLL-ENL fusion and a constitutively active N-RAS) independently of p53. Moreover, ATR inhibition as a single agent exhibited antitumoral activity, both reducing tumor burden after establishment and preventing tumors from growing, in an immunocompetent allograft mouse model of AMLMLL and in xenografts of a human AML-MLL cell line. We also found that inhibition of ATM, a kinase that senses DNA double-strand breaks, also promoted the survival of the AMLMLL mice. Collectively, these data indicated that ATR or ATM inhibition represent potential therapeutic strategies for the treatment of AML, especially MLL-driven leukemias.

INTRODUCTION

One recurrent finding in cancer is the presence of DNA replication stress, which, if persistent, leads to DNA double-strand breaks (DSBs) that initiate genomic rearrangements in cancer cells (1, 2). In addition to oncogenes, many of the agents used in genotoxic chemotherapy, including antifolates, nucleotide analogs, topoisomerase inhibitors, or platinum derivatives, are potent inducers of replication stress. In mammalian cells, the replication stress response (RSR) is a signaling cascade that initiates with the activation of the kinase ATR and activates the kinase checkpoint kinase 1 (CHK1; encoded by CHEK1) (3, 4). The presence of DNA replication stress renders cancer cells particularly dependent on a proficient ATR response for survival (5). Accordingly, studies with a hypomorphic ATR mouse model showed that low levels of ATR are particularly toxic for cells carrying oncogenic mutations (6, 7). Additionally, ATR or CHK1 inhibitors have exhibited efficacy in hematopoietic tumors, mostly in in vitro settings (611).

Although persistent replication stress or DNA damage activates the tumor suppressor p53, encoded by TP53, which triggers cell cycle arrest and apoptosis (12), cell death caused by low ATR activity is not only p53-independent but also enhanced by p53 deficiency (13, 14). Likewise, the toxicity of chemical inhibitors of ATR is higher in cells lacking p53 (10, 15, 16). This p53-independent cell killing by ATR inhibitors is linked to their capacity to induce the accumulation of replication stress and premature mitotic entry, activities that are unrelated to p53 functions (17, 18). Hence, ATR inhibitors offer an alternative for the elimination of p53-deficient tumors.

Previous studies have established that an intact p53 network is a critical determinant of the effectiveness of chemotherapy in acute myeloid leukemia (AML) (19). MLL (mixed lineage leukemia) refers to chromosomal translocation products involving the gene KMT2A, which encodes histone-lysine N-methyltransferase 2 (20). In contrast to other oncogenic fusion proteins, cells from AML patients with MLL fusion proteins (AML-MLL) do not mount an effective p53 response and are therefore resistant to current genotoxic treatments (19). Consistent with this, MLL rearrangements and mutations in TP53 rarely occur together in human AML (21, 22). Thus, alternative therapies are needed to overcome chemotherapy resistance associated with p53 dysfunction in AML-MLL. In addition to the need of a therapy that works on p53-deficient tumors, several lines of evidence suggested that targeting ATR could be particularly beneficial in AML-MLL. First, reduced amounts of ATR in mouse models inhibited growth of AML driven by the MLL-ENL oncogene, which encodes a fusion of KMT2A and the transcription activator ENL (7). Second, inhibitors of ATR or its target CHK1 are toxic to human cells and mouse models of several lymphomas and leukemias, including p53-deficient tumors (69, 11, 23). Moreover, the particular efficacy of RSR inhibitors in lymphoid tumors is consistent with a preferential role for the RSR in the untransformed lymphoid compartment, exemplified by the frequent presence of anemia in mice suffering from replication stress (13, 2427). Finally, inhibition of ATR or inhibition of the related DNA damage response (DDR) kinase ATM predisposed primary stem cells infected with retroviruses expressing MLL-AF9, a fusion between KMT2A and the transcription activator AF9, toward differentiation in vitro (28). On the basis of these data, we predicted that inhibition of ATR or ATM could have potential as a therapy for MLL-associated leukemia. Here, we report the antitumoral effects of ATR or ATM inhibitor monotherapy in an immunocompetent mouse model of AMLMLL.

RESULTS

Lymphomas and leukemias have high levels of CHEK1 expression

Because ATR inhibitors are preferentially toxic for cells experiencing replication stress, tumors with high endogenous levels of this stress could be promising targets. We previously showed that increased CHK1 reduces replication stress and improves the survival of cells expressing oncogenes or reprogramming factors (29, 30). On this basis, we predicted that high CHEK1 expression could be a signature of tumors with high levels of replication stress and, therefore, sensitivity to ATR inhibitors (29). This hypothesis is supported by work that identified an enrichment of CHEK1 gene amplifications in genomically unstable ovarian cancers, which correlated with an enhanced sensitivity to ATR inhibitors (31). Hence, CHK1 abundance may provide a biomarker for ATRi sensitivity.

To compare the relative levels of CHEK1 gene expression across different cancer types, we analyzed data from the human Cancer Cell Line Encyclopedia (32). CHEK1 mRNA was most abundant in Burkitt lymphomas, which we previously showed to be highly dependent on ATR and CHK1 for their survival (fig. S1) (6). In addition to Burkitt lymphoma, CHEK1 expression was distinctively high in various lymphomas and leukemias (fig. S1). Moreover, studies with hematopoietic tumor cells have shown good efficacy of ATR or CHK1 inhibitors (611). Overall, these analyses identify distinctively high levels of CHEK1 expression in lymphomas and leukemias, which could be indicative of underlying DNA replication stress. Consistent with our findings, in the companion paper in this issue, David et al. (33) report that CHEK1 expression and CHK1 abundance negatively correlate with prognosis in patients of AML.

AMLMLL cells in culture are highly sensitive to ATR inhibitors

From the different kinds of hematopoietic malignancies, we decided to focus on AML carrying MLL translocations for the following reasons. First, mouse genetic studies indicated that ATR abundance is particularly important for the viability of AML cells (7). Second, ATR-dependent phosphorylation of MLL is involved in the response to replication stress (34). Third, in vitro experiments showed that ATR inhibitors promote the differentiation of primary stem cells infected with MLL-AF9 (28). Finally, because of the p53-independent mechanism of cell killing by ATR inhibitors (18), they could potentially overcome the limitation impinged by p53-deficient responses in MLL-driven AML.

To test the efficacy of ATR inhibition in the treatment of AML in vivo, we used a previously described mouse cell line generated by transforming bone marrow cells with viruses expressing MLL-ENL and N-RASG12D (AMLMLL). These cells have an activating mutation in NRAS, which is also common in human AML-MLL patients (19), and these cells recapitulate the deficient p53 signaling and poor responses to conventional chemotherapy that are observed in the clinic with AML-MLL patients (19). A 1-day treatment of AMLMLL cells with the ATR inhibitor AZ20 (35) (hereafter referred to as ATRi) eliminated 75% of the viable cells in culture (Fig. 1A). The cytotoxicity of ATR inhibitors is mediated by forcing premature mitotic entry from G2, the phase during which DSBs are generated (18). Consistently, exposure to ATRi led to the disappearance of most AMLMLL cells from the G2 phase of the cell cycle (Fig. 1B). ATRi also resulted in the accumulation of DSBs, as indicated by the phosphorylation of the DDR targets KRAB-associated protein 1 (KAP1), structural maintenance of chromosomes 1 (SMC1), and histone H2AX (Fig. 1C). Moreover, depletion of p53 with a retrovirus expressing a p53-targeting short hairpin RNA (shRNA) did not rescue the cells from the toxic effects of ATRi (Fig. 1, D to F).

Fig. 1 p53-independent toxicity of ATRi in AMLMLL cells in culture.

(A) Fluorescence activated cell sorting (FACS) analysis showing the percentage of viable AMLMLL cells [identified by size and 4′,6-diamidino-2-phenylindole (DAPI) exclusion] either untreated or exposed to ATRi (10 μM, 24 hours); FSC-A, forward scattered light. Data are representative of two independent experiments. (B) FACS analysis of DNA content [propidium iodide (PI)] from the cultures used in (A) illustrating the depletion of G2 cells observed in response to ATRi. Data are representative of two independent experiments. (C) Western blot of SMC1, KAP1, and γH2AX phosphorylation and poly(ADP-ribose) polymerase 1 (PARP1) cleavage products in AMLMLL cells exposed to ATRi (5 μM, 6 hours). Data are representative of two independent experiments. (D) Western blot confirming the depletion of p53 in AMLMLL cells after infection with retroviruses expressing a p53-targeting shRNA. β-Actin abundance is shown as a loading control. (E) FACS analysis showing the percentage of viable AMLMLL cells (identified by size and DAPI exclusion) from control or p53 shRNA–infected AMLMLL cells either untreated or exposed to ATRi (10 μM, 24 hours). Data are representative of two independent experiments. (F) FACS analysis of DNA content (PI) from the cultures used in (E) illustrating the depletion of G2 cells observed in response to ATRi. Data are representative of two independent experiments.

To determine the mechanism for the sensitivity of AMLMLL cells to ATRi, we analyzed the RSR in these cells. Exposure to the ribonucleotide reductase inhibitor hydroxyurea (HU) stimulated the phosphorylation of the ATR targets CHK1 and replication protein A (RPA), ruling out a deficiency in activation and signaling through the replication stress pathway in these cells (Fig. 2A). We also compared the response to ATRi in AMLMLL cells in an equivalent cell line that was generated by transforming bone marrow cells with viruses expressing an AML1-ETO translocation (AMLETO). This models a p53-proficient type of AML that has better prognosis than AML patients with the MLL-ENL translocation (19). ATRi was more toxic for AMLMLL than for AMLETO cells (Fig. 2B). The enhanced sensitivity of AMLMLL cells correlated with higher levels of DNA damage in replicating cells, as measured by monitoring γH2AX phosphorylation of cells labeled with propidium iodide by FACS (Fig. 2C). ATRi induced a greater γH2AX signal, indicating a greater amount of damage, in AMLMLL cells. A structurally different ATR inhibitor, ETP-46464 (16), induced similar toxicity and accumulation of DNA damage (fig. S2). Furthermore, both ATR inhibitors induced cleavage of PARP1, an indicator of apoptosis (36), which confirms the toxicity of these inhibitors under conditions that induced DNA damage (fig. S2). Consistent with FACS data, AMLMLL and AMLETO cells exhibited differences in the rate of DNA replication fork progression when analyzed by stretched DNA fiber analyses. Whereas replication fork progression in AMLMLL cells was slower under basal conditions than that in AMLETO cells (Fig. 2D), exposure to ATRi had a bigger effect in reducing replication fork rates in AMLMLL cells, leading to an almost complete impairment of replication fork progression in these cells (Fig. 2D). In summary, MLL-driven AML cells exhibited an intrinsically higher sensitivity to ATR inhibitors than do AMLETO cells, with ATRi inducing the accumulation of replicative DNA damage, activation of the DDR, and p53-independent death in these cells.

Fig. 2 Increased levels of ATRi-induced replication stress in AMLMLL cells.

(A) Western blot of CHK1, RPA, and γH2AX phosphorylation in AMLMLL cells exposed to HU (2 mM, 2 hours). Data are representative of two independent experiments. (B) FACS analysis showing the percentage of viable AMLMLL and AMLETO cells (identified by size and DAPI exclusion) either untreated or exposed to ATRi (3 μM, 16 hours). Data are representative of two independent experiments. (C) FACS analysis of DNA content (PI) and H2AX phosphorylation in AMLMLL and AMLETO cells exposed to ATRi (10 μM, 5 hours). Data are representative of two independent experiments. (D) Fork rates were measured in stretched DNA fibers prepared from AMLMLL and AMLETO cells exposed (or not) to ATRi (10 μM, 5 hours). At least 200 tracks were measured per condition. ***P < 0.001 by two-tailed t test.

ATR inhibitors show efficacy as single agents in a mouse model of MLL-driven AML

To examine the in vivo efficacy of ATRi, we injected 105 AMLMLL tumor cells, which also expressed both green fluorescent protein (GFP) and luciferase for tracking, into immunocompetent mice (C57BL/6/BrdCrHsd-Tyrc). Even in immunocompetent recipients, intravenous injection of AMLMLL cells results in a very aggressive form of AML that infiltrates multiple organs and kills mice in a few weeks (19). In contrast to previous studies (19, 37), we did not irradiate recipient animals before the transplant. Irradiation depletes the bone marrow of the recipient mice, facilitating the expansion of the transplanted tumor. Because the tumor cells proliferated even in the absence of irradiation, we preferred to avoid this treatment to further mimic the normal context of AML. Transplanted mice were treated daily through an oral gavage with a dose of 60 mg/kg of ATRi, and tumor development was followed by monitoring luciferase activity with an in vivo imaging system (IVIS). Finally, to test the efficacy of ATR inhibitors, we used two protocols. In the prevention protocol, mice started receiving treatment on the day of the injection of AMLMLL cells (ATRiPr); in the therapy protocol, mice started receiving treatment after tumors were detectable by IVIS (ATRiTh).

Without drug treatment, AMLMLL cells rapidly expanded, leading to a lethal disease in control animals with a median survival of 23 days (Fig. 3A). The treatment of the ATRiTh group started at day 13. Both therapy and prevention groups showed a marked response as measured by IVIS at day 18 (Fig. 3B). IVIS on isolated organs confirmed that the treatment with ATRi severely limited tumor infiltration to organs, including liver, spleen, and lung (Fig. 3C). The decreased tumor burden was also evident from visual analysis of spleen sizes (Fig. 3D). Moreover, and in agreement with the observations made in vitro by exposure of cells to ATR inhibitors (Fig. 2C), treatment with ATRi led to a widespread accumulation of γH2AX-positive cells in the spleens of the treated mice (Fig. 3E). On day 23, we isolated bone marrow from mice of all groups and measured the presence of tumor cells by detection of GFP (Fig. 3F). GFP-positive cells were undetectable in the prevention group, and the therapy group exhibited a 17-fold decrease in the percentage of GFP-positive AMLMLL cells in the bone marrow. Consistent with this, videos of animals recorded on day 25 showed a clear improvement in the overall health of both ATRi-treated groups (movies S1 to S3). Although all animals eventually succumbed to leukemia, both groups showed a significant increase in the median life span (vehicle, 23 days; ATRiTh, 33 days; ATRiPR, 45 days) (Fig. 3A). Strikingly, at 40 days, all animals from the prevention group were alive, a time at which we decided to stop the treatment to explore potential curative effects of the therapy. Forty percent of these mice survived for more than 50 days, and one was alive for 117 days before succumbing to the disease.

Fig. 3 In vivo responses of AMLMLL to ATRi.

(A) Kaplan-Meier curves of AMLMLL transplanted mice that were either treated with vehicle (n = 9), ATRi from day 1 (ATRiPr; n = 7), or ATRi from day 13 (ATRiTh; n = 7). Treatment on the prevention group stopped at day 40. P value was calculated with the Mantel-Cox log-rank test. ***P < 0.001. (B) Representative IVIS of the luciferase signals observed on mice from the groups indicated on (A) on day 18. (C) Representative examples of the luciferase signal observed by IVIS on isolated organs from the indicated groups at day 23. (D) Picture of the spleen sizes observed at day 23 of the in vivo treatment experiment. Scale bar, 1 cm. (E) Representative images of γH2AX immunohistochemistry on spleens of AMLMLL transplanted mice treated with vehicle or ATRi (60 mg/kg, 11 days). Scale bar (black), 50 μm. Numbers indicate the percentage of γH2AX-positive cells in each case (means ± SD). (F) FACS analysis from the bone marrow collected from mice at day 23 of the treatment experiment indicated in (A). GFP (x axis) is used to monitor the presence of AMLMLL cells. y axis indicates FSC-A. The percentage of live GFP+ cells detected in each case is indicated. Data are representative of two independent groups. (G) Effect of ATRi as monotherapy on the growth of xenografts from the human MV4:11 cell line of MLL-driven AML. Treatment started when tumors became palpable, and eight animals were used per group. ***P < 0.001 by two-way analysis of variance.

Finally, we measured the efficacy of ATRi in xenografts of the human AML cell line MV4:11, which is also driven by an MLL-AF4 translocation (38). We subcutaneously implanted MV4:11 cells into the flanks of severe combined immunodeficient (SCID) mice. Treatment with ATRi started when tumors became palpable and was administered daily through oral gavage at 60 mg/kg. ATRi therapy significantly limited the growth of MV4:11 xenografts (Fig. 3G). In summary, our data showed that ATR inhibition elicits antitumoral responses when used as a single agent in allografts of mouse AMLMLL cells and in xenografts of a human AML cell line and provide an example of antitumor activity of this class of drugs in an immunocompetent model of cancer.

ATM inhibitors extend the survival of leukemia-bearing mice

Persistent replication stress leads to the breakage of replication forks and thus to DSBs that triggers an ATM-dependent DDR, suggesting an active role of ATM in limiting the toxicity of replication stress. Consistently, ATM deficiency is lethal in ATR-Seckel mice that accumulate high levels of replication stress (13). Moreover, previous in vitro data revealed that treatment of primary MLL-AF9 transformed cells with an ATM inhibitor or transformation of ATM knockout cells with MLL-AF9 results in poor growth and increased differentiation of these leukemic cells (28). To determine whether ATM is required for MLL leukemia in vivo, we first generated wild-type and ATM−/− AMLMLL tumors by transforming bone marrow hematopoietic progenitor cells of both genotypes retroviruses expressing MLL-AF9-IRES-neo and N-RASG12D-IRES-GFP. When injected into immunodeficient NRG (Rag1null;IL2rgnull) recipient mice, both ATM wild-type and ATM−/− AMLMLL cells caused lethal leukemia with no difference in median survival (Fig. 4A). Thus, even though loss of ATM activity inhibits growth in vitro (28), we found that it did not have a detectable impact on the development of MLL-AF9 leukemia in vivo in primary transplants.

Fig. 4 In vivo responses of AMLMLL to ATMi.

(A) Kaplan-Meier curves of mice that were transplanted with ATM wild-type (WT) (n = 6) and ATM−/− (n = 8) bone marrow hematopoietic progenitors that had been infected with retroviruses expressing MLL-AF9-IRES-neo and N-RASG12D-IRES-GFP. P value was calculated with the Mantel-Cox log-rank test. (B) Chemical structure of the ATM inhibitor AZD0156 (ATMi). (C) Representative IVIS of the luciferase signals observed on AMLMLL transplanted mice that were either treated with vehicle, ATMi from day 1 (ATMiPr), or ATMi from day 8 (ATMiTh). IVIS imaging was conducted on day 22 of the experiment. (D) Image of the spleen sizes observed at day 22 of the in vivo treatment experiment explained in (C). Scale bar, 1 cm. (E) Representative examples of the luciferase signal observed by IVIS on isolated organs from the indicated groups at day 22. (F) Kaplan-Meier curves of AMLMLL transplanted mice that were either treated with vehicle (n = 9), ATMi from day 1 (ATMiPr; n = 9), or ATMi from day 8 (ATMiTh; n = 9). Treatment on both groups stopped at day 40. P value was calculated with the Mantel-Cox log-rank test. ***P < 0.001.

Despite the absence of an impact of ATM deletion, lack of the kinase activity of ATM is not the same as loss of ATM. For example, ATM kinase–inactivating mutations lead to embryonic lethality in mice, whereas ATM knockout mice are viable and show less genome instability than kinase-dead mutants (39, 40). To determine the effects of ATM inhibition on MLL leukemia, we used the newly developed ATM inhibitor AZD0156 (referred to as ATMi hereafter) (41) (Fig. 4B).

To determine the efficacy of the ATMi, we repeated the same prevention and therapy protocols that we used to evaluate the ATR inhibitors. We injected AMLMLL cells through the tail vein into the mice and treated the mice daily with 20 mg/kg of the ATMi or with the vehicle. In the prevention protocol, treatment started on the day of injection (ATMiPr); in the therapy protocol, treatment started when tumors were first detectable by IVIS (ATMiTh). The untreated cohort behaved similarly to the previously shown experiments, with a median survival of 26 days. As with ATRi, the therapy with ATMi had a notable effect with either protocol, leading to reduced overall luciferase signal measured by IVIS (Fig. 4C), smaller spleen size (Fig. 4D), reduced organ infiltration (Fig. 4E), and prolonged survival of AMLMLL-injected mice (vehicle, 23 days; ATMiTh, 50 days; ATMiPR, 66 days) (Fig. 4F). Videos of mice recorded on day 23 confirmed the improvement in the overall health of ATMi-treated mice (movies S4 to S6).

DISCUSSION

Current treatment of acute pediatric leukemias involves the use of broad-spectrum genotoxic approaches. However, these lines of chemotherapy are largely ineffective for treatment of leukemias that have MLL translocations. In addition, some of these therapies for AML, such as the topoisomerase II inhibitor etoposide, counterproductively promote MLL translocations and therapy-related leukemia (42, 43). Thus, treatment of this subset of leukemias would benefit from a targeted therapy that exploits a specific vulnerability in the cancer cells.

One such vulnerability is associated with the function of KMT2A, which is the protein encoded in a gene associated with MLL translocation events. KMT2A is a lysine methyltransferase that functions as an epigenetic regulator (20). Leukemias carrying MLL fusion proteins require few, if any, additional mutations. Rather, fusion proteins induce leukemia by deregulating transcription at MLL fusion protein target genes, such as the HOXA gene cluster and MEIS1 (44, 45). Abnormal expression of these genes is associated with epigenetic changes, including alteration in DNA and histone methylation. For example, the histone 3 Lys79 (H3K79) methylase DOT1L is recruited to MLL fusion protein target genes, and this subtype of leukemia depends on DOT1L enzymatic activity (46). Similarly, MLL leukemias depend on several hematopoietic transcription factors, such as the bromodomain and extraterminal (BET) protein BRD4, to maintain their leukemic stem cell properties (31, 37, 47). Thus, one therapeutic approach for leukemogenesis resulting from MLL translocation events is to disrupt the MLL target gene expression program with drugs that target epigenetic-modifying enzymes or the products of genes that depend on such modifications (44). In this context, drugs that inhibit BET, such as the drug JQ1, or DOT1L are currently under clinical investigation.

In addition to lineage-specific transcriptional circuits, our experiments suggested that a second point of vulnerability in MLL-driven AML is the RSR. Whereas we found that the amount of replication stress markers (γH2AX) is not distinctively high in these tumors, we propose that this is because these cells have increased levels of RSR factors, such as CHK1, which help buffer the levels of replication stress. Accordingly, the accompanying paper from David et al. (33) reveals increased CHK1 levels in cells from human AMLMLL patients. We propose that, whereas this increase in RSR factors limits the basal toxicity of replication stress and thus facilitates tumor growth, it also represents a vulnerability because the tumor cells become particularly dependent on a proficient RSR. To what extent MLL translocations are responsible for the replication stress in AMLMLL cells, which have other defects such as N-RAS hyperactivity, remains to be established. Despite the hope for new targeted therapies, mechanisms for resistance to inhibitors of BET (31, 48, 49) or ATR (18) have been uncovered. Noteworthy, perturbing the chromatin-related functions of MLL fusion proteins leads to replication stress (34, 50). In this context, a combination of RSR inhibitors, leading to p53-independent toxicity, and epigenetic inhibitors, which may both interfere with the transcriptional properties of the MLL fusion protein and further increase the levels of replication stress, could help overcome the resistance of MLL-driven AML to chemotherapy.

MATERIALS AND METHODS

Cells and reagents

Mouse AML cells carrying the MLL-ENL translocation (plus IRES-GFP) and oncogenic N-RAS (luciferase-IRES-N-RASG12D), referred to in the text as AMLMLL, were developed as previously described (19). The retroviral plasmid with a p53-targeting shRNA was provided by M. Barbacid and transduced using standard protocols. MV4:11 cells were obtained from the American Type Culture Collection. Cells were cultured under standard conditions (5% CO2 and 20% O2) in RPMI-1640 (EuroClone) medium supplemented with 10% fetal bovine serum (FBS; Sigma-Aldrich) and penicillin/streptomycin (Pen/Strep; Life Technologies) (10 μg/ml). ATRi (AZ20, synthesized by GVK BIO), ATMi (AZD0156, AstraZeneca), and HU (Sigma-Aldrich) were used as indicated.

Flow cytometry

To measure viability, cells were collected, washed once with phosphate-buffered saline (PBS) (pH 7.4), stained in a DAPI solution [DAPI (0.2 μg/ml) in PBS], and analyzed by flow cytometry in a FACSCanto II (Becton-Dickinson) machine. For cell cycle profiles, cells were collected, washed with PBS, and fixed in suspension in ice-cold 70% (v/v) ethanol in PBS. After washing in PBS, cells were stained in PBS containing propidium iodide (10 μg/ml) and ribonuclease A (0.5 mg/ml) and collected in a Becton-Dickinson FACSCalibur machine. For DNA content and γH2AX analysis, p-Ser139 H2AX (Millipore) antibodies were used as previously described (18). Data were analyzed by FACSDiva (BD Biosciences) and FlowJo (Tree Star) softwares.

Protein extracts and Western blotting

For total protein extract preparation, AMLMLL cells were collected upon treatment with ATRi (5 μM for 6 hours) or HU (2 mM for 2 hours), washed once with PBS, and lysed in urea buffer [8 M urea, 1% CHAPS, and 50 mM tris-HCl (pH 8.0)] for 30 min at 4°C with agitation. Samples were resolved by SDS-polyacrylamide gel electrophoresis and analyzed by standard Western blotting techniques. The following primary antibodies were used: p-Ser345 Chk1 (Cell Signaling), p-Ser4/Ser8 RPA32 (Bethyl), p-Ser139 H2AX (Millipore), H2AX (Abcam), p-SMC1 [Monoclonal Antibody Unit, Spanish National Cancer Centre (CNIO)], p-Ser824 KAP-1 (Bethyl), and PARP1 (Cell Signaling). Alexa Fluor 680– or Alexa Fluor 800–conjugated secondary antibodies (Life Technologies) were used for detection with a LI-COR Odyssey infrared imaging system (LI-COR Biosciences).

Transplantation and in vivo treatment studies

For tumor induction and treatment studies, 105 AMLMLL cells were transplanted by tail vein injection into 8- to 12-week-old immunocompetent albino recipient mice (C57BL/6/BrdCrHsd-Tyrc). In the case of xenografts, 1.4 × 105 MV4:11 cells were injected subcutaneously into the flanks of SCID mice, and tumor growth was measured with a caliper. The mice were treated daily with ATRi (50 mg/kg; dissolved in 10% N-methyl-2-pyrrolidone/90% polyethylene glycol 300) or ATMi [20 mg/kg; dissolved in 10% dimethyl sulfoxide/90% Captisol (30%)], and the corresponding vehicles by oral gavage. Health status of the treated mice was monitored daily. The mice were maintained at the CNIO under standard housing conditions with free access to chow diet and water, in recommendation of the Federation of European Laboratory Animal Science Association. All mouse work was performed in accordance with the Guidelines for Humane Endpoints for Animals Used in Biomedical Research and under the supervision of the Ethics Committee for Animal Research of the Instituto de Salud Carlos III.

Monitoring of leukemias

To monitor tumor formation, AMLMLL transplanted mice were monitored every 3 to 4 days, starting 5 days after the injection of tumor cells, by bioluminescent imaging with an IVIS spectrum imaging system. The mice were intraperitoneally injected with 150 mg/kg of d-luciferin (Perkin-Elmer), anesthetized with isoflurane, and imaged for 30 s after 5 min after luciferin injection. For bioluminescence analysis of organs, the mice were injected with d-luciferin and euthanized by CO2, and the organs were collected and imaged for 5 s. To measure the persistence of AMLMLL cells, we isolated bone marrow from vehicle- or ATRi-treated animals by flushing femurs and tibias [RPMI-1640 medium supplemented with 10% FBS and Pen/Strep (10 μg/ml)]. Erythrocyte lysis was performed by treating bone marrow cells with a commercial ACK (ammonium-chloride-potassium) red lysis buffer (Lonza) for 5 min at room temperature. Cells were stained with c-KIT APC-H7 antibody (BD Biosciences) and analyzed in FACSCanto II (BD Biosciences; FACSDiva software). Data were analyzed with FlowJo (Tree Star) software.

Histopathology

Spleens were collected from vehicle- and ATRi-treated mice, fixed in formalin, and embedded in paraffin/formalin blocks. Sections and immunohistochemistry staining against γH2AX were performed following standard procedures. Slides were digitalized with Mirax Scan (Zeiss), and γH2AX-positive cells were automatically quantified from digitalized slides with AxioVision 4.6.3 software (Zeiss).

DNA fiber analyses

AML and AMLETO cells were pulse-labeled with 50 μM chlorodeoxyuridine (CldU) (20 min) followed by 250 μM iododeoxyuridine (IdU) (20 min). Labeled cells were collected, and DNA fibers were spread in buffer containing 0.5% SDS, 200 mM tris (pH 7.4), and 50 mM EDTA. For immunodetection of labeled tracks, fibers were incubated with primary antibodies [for CldU, rat anti-BrdU (bromodeoxyuridine); for IdU, mouse anti-BrdU] for 1 hour at room temperature and developed with the corresponding secondary antibodies for 30 min at room temperature. Mouse anti–single-stranded DNA antibody was used to assess fiber integrity. Slides were examined with a Leica DM6000 B microscope, as described previously (51). The conversion factor used was 1 μm = 2.59 kb (52).

SUPPLEMENTARY MATERIALS

www.sciencesignaling.org/cgi/content/full/9/445/ra91/DC1

Fig. S1. CHEK1 mRNA expression is highest in lymphomas and leukemias.

Fig. S2. Toxicity, replication stress, and DNA breakage induced by two distinct ATR inhibitors in AMLMLL cells.

Movie S1. Control mice for the ATRi experiment with animals injected with AMLMLL cells after treatment with vehicle.

Movie S2. Mice injected with AMLMLL cells and treated with ATRi on the prevention protocol.

Movie S3. Mice injected with AMLMLL cells and treated with ATRi on the therapy protocol.

Movie S4. Control mice for the ATMi experiment with animals injected with AMLMLL cells after treatment with vehicle.

Movie S5. Mice injected with AMLMLL cells and treated with ATMi on the prevention protocol.

Movie S6. Mice injected with AMLMLL and treated with ATMi on the therapy protocol.

REFERENCES AND NOTES

Acknowledgments: We thank J. Zuber for the reagents and discussions, A. L. Kung for help with human AML cells, and M. Barbacid for providing the plasmid containing the p53-targeting shRNA. Funding: Work in O.F.-C. laboratory was supported by Fundación Botín, by Banco Santander through its Santander Universities Global Division, and by grants from the Spanish Ministry of Economy and Competitiveness (MINECO) (SAF2014-57791-REDC and SVP-2013-068072), Fundació La Marato de TV3, the Howard Hughes Medical Institute, and the European Research Council (ERC-617840). The A.N. laboratory was supported by the Intramural Research Program of the NIH, the National Cancer Institute (NCI), the Center for Cancer Research, an Ellison Medical Foundation Senior Scholar in Aging, and the Alex Lemonade Stand Foundation Award. Author contributions: I.M.-P, A.D., M.M., V.L., M.E.A., A.T., H.-T.C., A.E., R.A., A.B., C.H., and X.W. contributed to the experiments; K.G.P., B.B., E.C., A.D., C.H., and A.J.P. worked in defining the experimental conditions for ATM and ATR inhibitors; S.A.A. helped in the analysis of the in vivo data; and O.F.-C. and A.N. designed the project and wrote the paper. Competing interests: AZD0156 was provided by AstraZeneca to the investigators under terms of a Materials Cooperative Research and Development agreement with the NCI. This compound has been patented by AstraZeneca. Data and materials availability: AZD0156 is available to third party investigators by AstraZeneca through an Open Innovation Portal request (http://openinnovation.astrazeneca.com/).
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