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ATR kinase inhibition sensitizes quiescent human cells to the lethal effects of cisplatin but increases mutagenesis
Mutat Res Fund Mol Mech Mutagen 816-818 (2019) 111678
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ATR kinase inhibition sensitizes quiescent human cells to the lethal effects of cisplatin but increases mutagenesis
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Rebekah J. Hutcherson, Michael G. Kemp
Department of Pharmacology and Toxicology, Wright State University Boonshoft School of Medicine, 3640 Colonel Glenn Hwy, Dayton, OH, 45435, United States
A R T I C LE I N FO
A B S T R A C T
Keywords: Cisplatin Chemotherapy DNA damage response Quiescence Translesion synthesis Protein kinase signaling
The ATR protein kinase is known to protect cells from DNA damage induced during the replicative phase of the cell cycle. Small molecule ATR kinase inhibitors have therefore been developed to improve the effectiveness of DNA damage-based chemotherapy regimens aimed at killing rapidly proliferating tumor cells. However, whether ATR functions in a similar manner in non-replicating cells has not been examined and is important considering the fact that most cells in the body, including cancer stem cells in solid tumors, normally reside in either a quiescent or differentiated non-replicating state. Using cultured human cell lines maintained in a quiescent or slowly growing state in vitro, ATR was found to be activated following treatment with the common anti-cancer drug cisplatin in a manner dependent on the nucleotide excision repair (NER) system. Moreover, treatment with the ATR kinase inhibitors VE-821 and AZD6738 enhanced quiescent cell killing and apoptotic signaling induced by cisplatin. However, ATR kinase inhibition in quiescent cells treated with a low concentration of cisplatin also elevated the level of mutagenesis at the hypoxanthine phosphoribosyltransferase locus and resulted in increased levels of PCNA mono-ubiquitination. These results suggest that the excision gaps generated by NER may require a greater utilization of potentially mutagenic translesion synthesis polymerases in the absence of ATR kinase function. Thus, though ATR kinase inhibitors can aid in the killing of cisplatin-treated quiescent cells, such treatments may also result in a greater reliance on alternative mutagenic DNA polymerases to complete the repair of cisplatin-DNA adducts.
1. Introduction DNA damaging compounds are routinely used in the treatment of a variety of different tumor types. Rapidly proliferating cancer cells are generally thought to be at greater susceptibility to the lethal effects of DNA damaging drugs than normal cells and cells that are not actively progressing through mitotic cell cycle [1]. Furthermore, the ability of such compounds to induce cell death may be limited by the cellular DNA damage response (DDR), which is composed of diverse biochemical systems and signaling pathways that promote cell survival and recovery through DNA repair, cell cycle checkpoints, and other pathways [2–4]. The Ser/Thr protein kinase ATR (ataxia telangiectasia and rad3related) is a major regulator of the DDR, particularly in cells undergoing chromosomal DNA replication [5]. ATR has therefore recently emerged as a novel target for cancer chemotherapy regimens that are aimed at
improving the effectiveness of commonly used agents that generate DNA damage and replication stress [6–8]. Using diverse model organisms and systems ranging from yeast to frog egg extracts to cultured human cells, a plethora of studies have demonstrated that ATR limits replicating cells from the lethal effects of DNA damage by stabilizing stalled replication forks, inhibiting new replication origin firing, delaying the entry of cells into mitosis, enabling translesion synthesis, and promoting DNA repair and recombination [5]. Because nearly all of these events are specific to cells in S phase, our understanding of ATR function in the DDR is largely restricted to cells that are actively synthesizing DNA and progressing through the mitotic cell cycle. Given that most cells in the body are in a non-replicating quiescent or differentiated state, it is important to understand whether ATR can become activated in non-replicating cells and how ATR signaling impacts cellular responses to DNA damage that occur independent of canonical chromosomal DNA replication. For example, a recent study
Abbreviations: ATR, ataxia telangiectasia-mutated and rad3-related; NER, nucleotide excision repair; DDR, DNA damage response; TLS, translesion synthesis; HPRT, hypoxanthine phosphoribosyltransferase; SP, spironolactone; XPB, xeroderma pigmentosum group B; NA-AAF, N-acetyoxy-2-acetylaminofluorene; ssDNA, singlestranded DNA; RPA, replication protein A; PCNA, proliferating cell nuclear antigen ⁎ Corresponding author. E-mail address: [email protected] (M.G. Kemp). https://doi.org/10.1016/j.mrfmmm.2019.111678 Received 26 June 2019; Received in revised form 19 August 2019; Accepted 15 September 2019 Available online 17 September 2019 0027-5107/ © 2019 Elsevier B.V. All rights reserved.
Mutat Res Fund Mol Mech Mutagen 816-818 (2019) 111678
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weeks as previously reported [11,12].
found that ATR inhibition can either promote cell death or survival in response to treatment with the bulky DNA adduct-inducing fluorene metabolite N-acetoxy-2-acetylaminofluorene (NA-AAF) depending on whether the cells are in a replicating/cycling or non-replicating/noncycling state [9,10], respectively. Whether these opposing functions for ATR are seen in response to commonly used anti-cancer drugs is not known. This lack of knowledge is a potential concern because ATR kinase inhibitors are entering clinical trials as adjuvants in cancer chemotherapy regimens. Thus, the toxicity and mutagenicity of chemotherapy drugs in non-replicating normal cells and tissues and quiescent cancer stem cells may be positively or negatively impacted by the addition of an ATR kinase inhibitor. Using cisplatin as a model anti-cancer drug, we investigated the impact of small molecule ATR inhibitor co-administration in non-replicating, quiescent human cells in vitro. We observed that ATR is capable of becoming activated in quiescent cells treated with cisplatin and that ATR kinase inhibition sensitizes quiescent cells to the lethal effects of cisplatin. Though this would be a favorable outcome in nonreplicating tumor cells in vivo, we also found that ATR inhibition increased the level of mutagenesis and resulted in increased monoubiquitination of PCNA, which may imply a greater reliance on the potentially mutagenic translesion synthesis (TLS) pathway to fill in the gaps generated by the NER machinery. Thus, ATR kinase inhibition may have both positive and negative effects on quiescent cell responses to DNA damaging compounds that are commonly used to treat human cancers.
2.3. Protein immunoblotting Cells were washed with cold PBS, scraped from the plate, and pelleted by gentle centrifugation. Cells were then lysed for 20 min on ice in 20 mM Tris−HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, and 1% Triton X-100. Following centrifugation in a microcentrifuge for 10–15 min at maximum speed, the soluble cell lysates were transferred to new tubes for immunoblot analysis. Chromatin-associated proteins were obtained from cells following two extractions with a modified cytoskeletal buffer (10 mM Tris−HCl (pH 7.4), 100 mM NaCl, 3 mM MgCl2, 1 mM EDTA, 1 mM Na3VO4, 10 mM NaF, and 0.1% Triton X100). Equal amounts of cell/chromatin extracts were separated by SDSPAGE, transferred to nitrocellulose, and then probed by immunoblotting using standard procedures. Primary antibodies included antibodies against ATR (sc-1887), actin (I-19), PCNA (sc-10) from Santa Cruz Biotechnology, phospho-p53 (Ser-15, 9284) from Cell Signaling Technology, phospho-ATR (Thr-1989, GTX128145) from GeneTex, and RPA70 (A300-421A) from Bethyl Laboratories. All primary antibodies were used at a 1:1000 or 1:2000 dilution in 1X TBST (50 mM Tris−HCl pH 7.4, 135 mM NaCl, and 0.1% Tween 20). Chemiluminescence was visualized with Clarity Western ECL substrate (Bio-Rad) or SuperSignal West Femto substrate (Thermo Scientific) using a Molecular Imager Chemi-Doc XRS + imaging system (Bio-Rad). Signals within the linear range of detection were quantified using Image Lab software (Bio-Rad) as previously described [9,10]. All experiments analyzing DNA damage response signaling were repeated at least two times, as indicated, and the average (and standard error) relative level of protein/phosphoprotein expression was determined and plotted.
2. Materials and methods 2.1. Cell culture Human immortalized HaCaT keratinocytes and U2OS osteosarcoma epithelial cells were cultured at 37 °C in a 5% CO2 humidified incubator in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% Fetal Clone III (Hyclone), 6 mM L-glutamine, 100 units/ml of penicillin, and 100 μg/ml of streptomycin. Cycling, replicating cells were typically plated at 30% confluence and treated the following day. Noncycling, quiescent cells were plated at 40–60% confluence, grown for 2 days in normal medium until the cells reached confluence, and then maintained for 2–3 days in DMEM containing 0.5% Fetal Clone III and penicillin/streptomycin before being subjected to experimental treatments. The ATR inhibitors VE-821 and AZD6738 (Selleckchem) were dissolved in DMSO and diluted 1000-fold to be used at the final concentrations of 10 μM and 3 μM, respectively. Spironolactone (SP; dissolved in DMSO) was used at a final concentration of 10 μM. Cisplatin (Sigma) was dissolved in PBS at a 3 mM stock concentration and then added to cell culture medium to obtain the indicated final concentrations. Cells were pre-treated for 2 h with SP and 30 min with the ATR inhibitors before addition of cisplatin to the culture medium. All compounds remained in the culture medium for all acute assays of survival, DNA repair, and kinase signaling.
2.4. DNA immunoblotting Bromodeoxyuridine (BrdU) was added to the culture medium at final concentration of 10 μg/ml for 15 min before harvesting the cells. Genomic DNA was purified using the GenElute Mammalian Genomic DNA Miniprep Kit (Sigma) and quantified using PicoGreen fluorescence (Invitrogen) on a Synergy H1 Hybrid Multi-Mode microplate reader (BioTek). DNA was immobilized on nitrocellulose, dried, and immunoblotted with antibodies against BrdU (Sigma B8434), cisplatinmodified DNA (CP9/19; Abcam ab103261), or single-stranded DNA (Millipore MAB3034) as previously described [10]. 3. Results 3.1. ATR kinase inhibition sensitizes replicating cells to cisplatin To confirm previous reports that ATR kinase inhibition sensitizes asynchronously growing human cells to the chemotherapeutic drug cisplatin, sub-confluent HaCaT keratinocytes and U2OS osteosarcoma cells in the logarithmic phase of growth were pre-treated with vehicle or the ATR inhibitor (ATRi) VE-821 for 30 min and then exposed to increasing concentrations of cisplatin. As shown in Fig. 1A, the inhibition of ATR potentiated the lethal effects of cisplatin treatment. These results demonstrate that ATR limits the effectiveness of cisplatin at killing proliferating HaCaT and U2OS cells. To compare the response of replicating cells to that of non-replicating, quiescent cells, sub-confluent proliferating HaCaT and U2OS cells and cells grown to confluence and maintained in medium with a low concentration of growth factors were pulsed with the nucleotide analog BrdU to measure the level of DNA synthesis by genomic DNA immunoblotting. As demonstrated in Fig. 1B, both the HaCaT and U2OS cells incorporated significantly reduced levels of BrdU when in the confluent, growth factor-deprived state. These results indicate that a quiescent or slowly growing cell state can be readily generated in vitro with these cell lines, and thus for simplicity we will refer here to both
2.2. Cell survival, cell death, and mutagenesis assays Crystal violet staining was used to monitor cell survival 3 days after cisplatin treatment as previously described [9,10]. Quantitation of trypan blue-positive cells was measured using a Countess II Automated Cell Counter (ThermoFisher). Measurements of sub-G1/G0 cells was carried out with an Accuri C6 flow cytometer following staining of ethanol-fixed cells with propidium iodide. For long-term assays of cell recovery and mutagenesis, drug-containing medium was replaced with fresh, low-serum medium after 2 days of treatment. For clonogenic survival assays, cells were then trypsinized 3 days later, re-plated at 100 cells per 100 mm plate, and then stained with crystal violet 10–14 days later to count surviving colonies. Cells with mutations at the HPRT locus were quantified by selection with 4 μM of 6-thioguanine for 2–3 2
Mutat Res Fund Mol Mech Mutagen 816-818 (2019) 111678
R.J. Hutcherson and M.G. Kemp
Fig. 1. Cisplatin induces cell death in both proliferating and quiescent human cells. Subconfluent, proliferating (A) HaCaT keratinocytes and (B) U2OS osteosarcoma cells were pre-treated with vehicle (DMSO) or the ATR inhibitor VE-821 (10 μM) for 30 min before treatment with the indicated concentrations of cisplatin. Surviving cells were stained with crystal violet 3 days later, and the stain was then solubilized and quantified using a spectrophotometer. Proliferating and quiescent (C) HaCaT and (D) U2OS cells were pulsed with BrdU for 15 min and then BrdU content in genomic DNA was measured via DNA immunoblotting to quantify the relative level of DNA synthesis. (E) HaCaT and (F) U2OS cells in either proliferating or quiescent state were exposed to increasing concentrations of cisplatin, and cell survival was measured 3 days later. The asterisks indicate significant differences (p < 0.05) in survival or DNA synthesis between treatment groups.
Immunoblotting of protein lysates from these cells revealed that both ATR itself [13,14] and the tumor suppressor protein p53 exhibited increased phosphorylation after cisplatin treatment (Fig. 2A), including at even lower concentrations of cisplatin (Supplementary Fig. 1). Pretreatment of the cells with two different small molecule inhibitors of ATR (VE-821 and AZD6738) largely blocked this response (Fig. 2A, B), which indicates that ATR is responsible for these phosphorylation events in quiescent cells. The canonical model for ATR kinase activation is that the stalling of DNA polymerases at DNA lesions results in the uncoupling of replicative helicase and polymerase activities [15], which subsequently leads to the generation of single-stranded DNA (ssDNA) that becomes coated with RPA to enable the recruitment and activation of ATR [16] in conjunction with the ATR-activating protein TopBP1 [5,17,18]. In
lines as being in a quiescent or non-replicating state. Non-replicating cells are generally considered to be much more resistant to DNA damaging agents than replicating cells [1]. Consistent with this notion, we observed that the non-replicating, quiescent HaCaT and U2OS cells required much higher concentrations of cisplatin to kill cells in comparison to their proliferating counterparts (Fig. 1C). 3.2. ATR kinase activity is stimulated in cisplatin-treated quiescent cells in a nucleotide excision repair-dependent manner We next examined whether the ATR kinase becomes activated in non-replicating cells treated with cisplatin. Quiescent HaCaT and U2OS cells were therefore exposed for 24 h to concentrations of cisplatin that had little effect on acute cell survival (60 μM and 30 μM, respectively). 3
Mutat Res Fund Mol Mech Mutagen 816-818 (2019) 111678
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Fig. 2. Cisplatin induces ATR kinase activation in an NER-dependent manner in quiescent human cells. (A) Quiescent HaCaT and U2OS cells were treated for 24 h with the indicated inhibitors and with cisplatin (60 μM or 30 μM, for HaCaT and U2OS, respectively), and then immunoblotting was performed to detect the activation of ATR kinase signaling. The asterisk indicates a non-specific band on the p53 immunoblot in U2OS cells. (B) Quantification of 3 independent experiments performed as described in (A). (C) Quiescent HaCaT cells were treated with DMSO or 10 μM spironolactone (SP) for 2 h before treatment for 24 h with 60 μM cisplatin. Protein lysates were analyzed by immunoblotting, and the results from 3 independent experiments were quantified and graphed. The asterisks indicate a significant difference (p < 0.05) in ATR autophosphorylation following cisplatin treatment.
sensitized the quiescent cells to killing by cisplatin (Fig. 3A). Similar results were obtained with the ATR inhibitor AZD6738 (Fig. 3B), in cells treated with the related platinating drug carboplatin (Fig. 3C), with trypan blue staining to quantify cells with a loss of membrane integrity (Supplementary Fig. 2), and in telomerase-immortalized normal human fibroblasts (data not shown). These results demonstrate that ATR kinase signaling is important for quiescent cells to properly respond to cisplatin treatment. Because previous studies revealed that ATR promoted the death of quiescent cells treated with the fluorene metabolite NA-AAF [9,10], the current data further suggest that though cisplatin and NA-AAF both induce the formation of DNA adducts that can be targeted for removal by the NER machinery, the inhibition of ATR kinase signaling can have dramatically different effects on cell fate. Cisplatin-induced cell death may occur via apoptosis, and thus we next examined apoptotic signaling in quiescent HaCaT and U2OS cells treated with cisplatin and an ATR inhibitor. Using the cleavage of the caspase substrate PARP as a measure of apoptotic signaling, we observed that treatment with the ATR kinase inhibitor VE-821 resulted in more PARP cleavage in quiescent HaCaT cells exposed to a range of concentrations of cisplatin than in DMSO-treated cells (Fig. 4A,B). Similar results were obtained with the ATR inhibitor AZD6738 (Fig. 4C). Analysis of DNA degradation by flow cytometric analysis of propidium
contrast, several studies have reported that a major pathway for ATR kinase activation in UV-irradiated quiescent cells occurs via single stranded DNA (ssDNA) intermediates that are generated as a result of UV photoproduct removal by the NER machinery [19–22]. To examine whether NER is required for ATR kinase activation in quiescent cells treated with cisplatin, non-replicating HaCaT cells were pre-treated with vehicle or with spironolactone (SP), which inhibits NER by causing the rapid proteolytic degradation of the core NER protein XPB [11,23]. As shown in Fig. 2C and Supplementary Fig. 1, pre-treatment with spironolactone largely blocked the cisplatin-dependent increase in ATR and p53 phosphorylation. We conclude that ATR can become activated in non-replicating, quiescent cells in a manner dependent on the NER machinery. 3.3. ATR kinase inhibition sensitizes cisplatin-treated quiescent cells to undergo an apoptotic form of cell death To examine the possible functions of ATR kinase signaling in cisplatin-treated quiescent cells, HaCaT and U2OS cells maintained in a non-replicating state were treated with the ATR inhibitor VE-821 and then exposed to increasing concentrations of cisplatin. Cell survival was then quantified 3 days later. These experiments revealed that VE-821 4
Mutat Res Fund Mol Mech Mutagen 816-818 (2019) 111678
R.J. Hutcherson and M.G. Kemp
Fig. 3. ATR kinase inhibition sensitizes quiescent human cells to cisplatin and carboplatin. (A) Quiescent HaCaT and U2OS cells were treated with vehicle (DMSO) or 10 μM VE-821 and the indicated concentration of cisplatin for 3 days, and then surviving cells were stained and quantified. (B) Experiments were performed as in (A) except that the ATR inhibitor AZD6738 (3 μM) was used. (C) Cells were treated as in (A) except that carboplatin was used in place of cisplatin. The asterisks indicate significant differences (p < 0.05) in survival between treatment groups at the concentrations of cisplatin or carboplatin.
iodide-stained cells further confirmed these findings (Fig. 4D) Thus, we conclude that ATR kinase inhibition enhances the ability of cisplatin to induce cell death via apoptosis in non-replicating quiescent cells.
3.4. ATR kinase inhibition results in increased mutagenesis in quiescent cells treated with cisplatin Though DNA damage can exert toxic effects on acute cell viability, the damage can also have potentially mutagenic and long-lasting effects
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Fig. 4. ATR kinase inhibition potentiates apoptotic signaling in cisplatin-treated quiescent cells. (A) Quiescent HaCaT cells were treated with DMSO or VE-821 and the indicated concentration of cisplatin for 48 h, after which cell lysates were immunoblotted to detect apoptotic signaling. (B) Quantification of results from at least three independent experiments performed as in (A). (C) Quiescent HaCaT cells were treated DMSO, VE-821 (10 μM) or AZD6738 (3 μM) and 60 μM cisplatin for 48 h. Cell lysates were probed with the indicated antibodies to monitor apoptotic signaling. (D) Cells treated for 4 days were stained with propidium iodide and analyzed by flow cytometry. The asterisks indicate significant differences (p < 0.05) in the percentage of cleaved PARP and sub-G1/G0 cells between the ATRi- and DMSO-treatment groups.
Fig. 5. ATR kinase inhibition in quiescent cells does not impact proliferative potential but results in increased mutagenesis. (A) Schematic of experiments examining how quiescent cells recover from treatment with cisplatin and an ATR kinase inhibitor. Quiescent HaCaT cells treated with a low concentration (10 μM) of cisplatin in the absence or presence of the ATR inhibitor VE-821 were re-plated 5 days later at low density in the presence of growth factors to allow for measurements of (B) clonogenic survival and (C) mutagenesis at the HPRT locus. The asterisk indicates a significant difference (p < 0.05) in the number of HPRT mutants as determined in three separate experiments.
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on cell proliferation. Moreover, quiescent cells are frequently induced to proliferate upon cell stress or tissue damage, and thus we next designed experiments to determine how ATR kinase inhibition in the nonreplicating, quiescent state impacts the ability of cisplatin-treated cells to recover and proliferate. A schematic of this experimental design is provided in Fig. 5A and involved the use of a low concentration of cisplatin that does not lead to detectable decreases in acute cell survival (Figs. 1 and 3). Interestingly, using a classical clonogenic survival assay to monitor cell recovery, we observed similar numbers of surviving colonies in the absence and presence of the ATR kinase inhibitor VE821 (Fig. 5B). Thus, ATR inhibition does not appear to significantly impact the long-term proliferative capacity of quiescent cells treated with a low concentration of cisplatin. In addition to inducing cell death, DNA adducts may also lead to mutagenesis. We therefore used the classical HPRT mutagenesis assay to quantify the number of cells that develop mutations at the HPRT locus, which enables cells to become resistant to the toxic nucleotide analog 6-thioguanine. As shown in Fig. 5C, ATR kinase inhibition resulted in an approximately 2.5-fold increase in the number of cisplatintreated cells with mutations at the HPRT locus. We therefore conclude that although ATR kinase inhibition in quiescent cells treated with a low concentration of cisplatin does not lead to significant changes in colony forming ability, the loss of ATR kinase activity under these conditions promotes mutagenesis. 3.5. ATR kinase inhibition does not impact the removal of cisplatin-DNA adducts by NER Cisplatin treatment results in the formation of intra-strand cisplatinDNA adducts that are potentially lethal and mutagenic to cells if not removed by the NER machinery. To determine whether the enhancement in cisplatin toxicity and mutagenicity by ATR inhibition in quiescent cells is correlated with a defect in cisplatin-DNA adduct removal, non-replicating HaCaT cells were treated with cisplatin in the presence or absence of the ATR inhibitor VE-821 and then genomic DNA was analyzed for cisplatin-DNA adduct content by immunoblotting. As shown in Fig. 6, the inhibition of ATR kinase activity did not significantly affect the rate of cisplatin-DNA adduct formation or removal from quiescent cell genomic DNA. These results indicate that the cisplatin lethality and mutagenicity that is exacerbated by ATR kinase inhibition in quiescent cells is not due to a defect in the damage removal step of NER.
Fig. 7. ATR kinase inhibition leads to increased PCNA monoubiquitination in cisplatin-treated quiescent cells. (A) Quiescent HaCaT cells were treated for 24 h with DMSO or VE-821 and the indicated concentration of cisplatin. Cells were then fractionated to enrich for chromatin-associated proteins, which were analyzed by immunoblotting with the indicated antibodies. The asterisk in the RPA70 blot indicates a non-specific band, and the characteristic monoubiquitinated form of PCNA is labeled as PCNA-ub. (B) Quantitation of results from 3 independent experiments performed as in (A). (C) Cells were pre-treated with spironolactone (SP) before exposure to cisplatin for 18 h and fractionation to enrich for the monoubiquitinated form of PCNA. The asterisks on the graphs indicate significant differences in RPA and mono-ubiquitinated PCNA chromatin association between the treatment groups (p < 0.05).
3.6. Inhibition of ATR results in elevated PCNA monoubiquitination Though cisplatin-DNA adduct removal by NER was not impacted by ATR inhibition (Fig. 6), earlier data indicating that ATR kinase activation is NER-dependent (Fig. 2C) suggests that a step of NER that occurs after damage excision may be relevant to the function of ATR in promoting survival and limiting mutagenesis. To examine this further,
Fig. 6. The rate of removal of cisplatin-DNA intrastrand adducts is not affected by ATR kinase inhibition. (A) Quiescent HaCaT cells were treated 30 μM cisplatin and either DMSO or VE-821 and then genomic DNA immunoblotting was performed at the indicated time points to detect the presence of cisplatin-DNA adducts. Blots were re-probed to detect total DNA (B) Quantitation of results from 2 independent experiments performed as in (A).
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that is observed at the HPRT locus in cisplatin-treated quiescent cells with inactive ATR (Fig. 5) may be due DNA synthesis by mutagenic DNA polymerases at NER gaps. Though this process promotes cell survival, the gap filling process may therefore be more mutagenic. A schematic summarizing these results is provided in Fig. 8.
quiescent HaCaT cells pre-treated with either vehicle or the ATR inhibitor VE-821 were exposed to cisplatin and then fractionated to enrich for chromatin-associated proteins. As shown in Fig. 7A, the levels of RPA and PCNA, which are known to associate with the ssDNA gaps generated by NER [24–26], were elevated on chromatin in a cisplatin dose-dependent manner. RPA is the major ssDNA-binding protein in human cells, and PCNA is a replicative polymerase clamp protein that facilitates gap filling DNA synthesis during NER [24,25,27]. Interestingly, we also observed the presence of a characteristic monoubiquitinated form of PCNA that was readily apparent in cisplatintreated quiescent cells. This modification is frequently used as a biochemical indicator for activation of the translesion synthesis (TLS) pathway of DNA synthesis [28], in which specialized polymerases are recruited to DNA via ubiquitin-binding domains to carry out DNA synthesis across damaged nucleotides or at difficult to replicate genomic regions. Thus, the TLS pathway is likely required in quiescent cells to fill at least a sub-set of excision gaps generated by the NER machinery [29]. Interestingly, though treatment with the ATR kinase inhibitor did not impact total PCNA levels on chromatin and resulted in only moderately higher levels of chromatin-associated RPA after treatment with a low dose of cisplatin (Fig. 7B), the levels of monoubiquitinated PCNA were 2- to 3-fold higher in cells treated with the ATR inhibitor. Treatment of cells with spironolactone to inhibit NER largely blocked PCNA monoubiquitination (Fig. 7C). Preliminary studies with quiescent HaCaT cells exposed to UVB radiation has revealed a similar phenotype upon ATR kinase inhibition (Shaj and Kemp, unpublished). Together, these results indicate that ATR inhibition may result in a greater dependence on the TLS pathway to complete gap filling DNA synthesis. However, additional work is needed to determine how the loss of ATR signaling impacts recruitment of specific DNA polymerases to excision gaps. Nonetheless, the increased mutagenesis
4. Discussion Cisplatin remains a widely used chemotherapeutic agent for the treatment of a variety of solid tumors. However, because the various DNA damage responses regulated by ATR, including DNA repair, cell cycle arrest, and replication fork stabilization may limit the effectiveness of cisplatin, clinical trials are currently underway to test whether ATR inhibition can potentiate the cancer cell killing effects of cisplatin and other DNA damaging drugs that generate replication stress [6,8]. Preclinical models using cancer cell lines in vitro [30–34] and tumor xenografts in mice in vivo [35–38] support the concept that ATR kinase inhibition can potentiate the ability of cisplatin to slow or prevent tumor growth. A great deal of knowledge has been gained on ATR function in the response to replication stress induced by cisplatin and other DNA damaging agents. Though several studies have demonstrated that DNA damaging agents can activate ATR kinase signaling in quiescent or G1 phase cells [19–21,39–45], the actual function of this signaling in nonreplicating quiescent cells is largely unexplored. This is an important issue because most cells in the body, including cancer stem cells, frequently reside in a non-replicating quiescent state. Cancer treatments that eliminate such cells may therefore be most effective at preventing tumor recurrence [46], and thus ATR inhibitors could aid in this process. However, cancer drugs can also induce mutations that drive tumor recurrence and carcinogenesis, and thus care must be taken to design Fig. 8. Schematic summarizing the effect of ATR kinase inhibition in cisplatin-treated quiescent cells. Under normal conditions, cisplatin-DNA adducts (indicated by G < > G) are removed by the NER machinery, ATR becomes activated, and the excision gap can be filled in by DNA synthesis and ligation. However, when ATR kinase is inhibited, the gap filling process becomes more dependent on the translesion synthesis (TLS) pathway, which can result in mutagenesis or cell death.
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treatment regimens that maximize cancer cell killing while also minimizing toxicity and mutagenicity in normal cells. Here we used an in vitro model of cell quiescence to better understand ATR kinase function. We found that ATR becomes activated in non-replicating quiescent cells in vitro in a manner dependent on the NER (Fig. 2) and that inhibiting ATR under these circumstances improves the cell killing and pro-apoptotic effects of cisplatin in non-replicating cells (Figs. 3 and 4). These results indicate that similar to the situation in replicating cells, ATR protects non-replicating cells from the damage caused by cisplatin. Interestingly, these results are in striking contrast to similar experiments in which replicating and non-replicating cells were treated with the fluorene metabolite N-acetoxy-2-acetylaminofluorene (NA-AAF) [9,10]. These previous studies showed that though ATR kinase inhibition sensitized replicating cells to NA-AAF, ATR inhibitors protected non-replicating cells from the lethal and apoptotic effects of NA-AAF. Though both cisplatin and NA-AAF generate DNA adducts that can be repaired by the NER system, it is possible that the bulkier adducts induced by NA-AAF are a stronger block to transcription than those induced by cisplatin, and thus transcription stalling-dependent activation of ATR may play a more prominent role in promoting apoptosis in that context [41]. It will therefore be important to examine how ATR kinase inhibitors modify cellular responses to DNA damage caused by different carcinogenic and anti-cancer compounds in non-replicating cells to better understand the signals that activate ATR as well as the downstream processes that are regulated by ATR. Nonetheless, our results here along with previous results using ionizing radiation [45] indicate that combination radio- and chemotherapies involving cisplatin and an ATR inhibitor may be useful to kill both the replicating and non-replicating cells that comprise solid tumors in vivo. However, such treatments could exacerbate the potentially toxic effects of cisplatin and other DNA damaging agents on specific organ systems. Moreover, our evidence that ATR kinase inhibition increases the mutation frequency and PCNA monoubiquitination (a signal for recruitment of TLS polymerases) (Figs. 5 and 7) suggests that anti-cancer therapies utilizing ATR kinase inhibitors may increase the risk of mutagenesis and carcinogenesis within quiescent cancer stem cells or in other tissues of the body. Thus, the risk of secondary tumors arising as a result of cisplatin-based anti-cancer therapies may be elevated by treatment with an ATR kinase inhibitor and should be monitored accordingly.
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Marti, E. Hefner, L. Feeney, V. Natale, J.E. Cleaver, H2AX phosphorylation within the G1 phase after UV irradiation depends on nucleotide excision repair and not DNA double-strand breaks, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 9891–9896. [20] F. Marini, T. Nardo, M. Giannattasio, M. Minuzzo, M. Stefanini, P. Plevani, M.M. Falconi, DNA nucleotide excision repair-dependent signaling to checkpoint activation, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 17325–17330. [21] S. Hanasoge, M. Ljungman, H2AX phosphorylation after UV irradiation is triggered by DNA repair intermediates and is mediated by the ATR kinase, Carcinogenesis 28 (2007) 2298–2304. [22] M. Matsumoto, K. Yaginuma, A. Igarashi, M. Imura, M. Hasegawa, K. Iwabuchi, T. Date, T. Mori, K. Ishizaki, K. Yamashita, M. Inobe, T. Matsunaga, Perturbed gapfilling synthesis in nucleotide excision repair causes histone H2AX phosphorylation in human quiescent cells, J. Cell. Sci. 120 (2007) 1104–1112. [23] S. Alekseev, M. Ayadi, L. Brino, J.M. Egly, A.K. Larsen, F. Coin, A small molecule screen identifies an inhibitor of DNA repair inducing the degradation of TFIIH and the chemosensitization of tumor cells to platinum, Chem. Biol. 21 (2014) 398–407. [24] R.D. Wood, Nucleotide excision repair in mammalian cells, J. Biol. Chem. 272 (1997) 23465–23468. [25] J.T. Reardon, A. Sancar, Nucleotide excision repair, Prog. Nucleic Acid Res. Mol. Biol. 79 (2005) 183–235. [26] A. Sancar, Mechanisms of DNA repair by photolyase and excision nuclease (nobel lecture), Angew. Chem. Int. Ed. Engl. 55 (2016) 8502–8527. [27] M.G. Kemp, J. Hu, PostExcision events in human nucleotide excision repair, Photochem. Photobiol. 93 (2017) 178–191. [28] J.E. Sale, Translesion DNA synthesis and mutagenesis in eukaryotes, Cold Spring Harb. Perspect. Biol. 5 (2013) a012708. [29] S. Sertic, A. Mollica, I. Campus, S. Roma, E. Tumini, A. Aguilera, M. Muzi-Falconi, Coordinated activity of Y family TLS polymerases and EXO1 protects non-S phase cells from UV-induced cytotoxic lesions, Mol. Cell 70 (2018) 189. [30] C.J. Huntoon, K.S. Flatten, A.E. Wahner Hendrickson, A.M. Huehls, S.L. Sutor, S.H. Kaufmann, L.M. Karnitz, ATR inhibition broadly sensitizes ovarian cancer cells to chemotherapy independent of BRCA status, Cancer Res. 73 (2013) 3683–3691. [31] H.J. Kim, A. Min, S.A. Im, H. Jang, K.H. Lee, A. Lau, M. Lee, S. Kim, Y. Yang, J. Kim, T.Y. Kim, D.Y. Oh, J. Brown, M.J. O’Connor, Y.J. Bang, Anti-tumor activity of the ATR inhibitor AZD6738 in HER2 positive breast cancer cells, Int. J. Cancer 140 (2017) 109–119. [32] K.B. Leszczynska, G. Dobrynin, R.E. Leslie, J. Ient, A.J. Boumelha, J.M. Senra, M.A. Hawkins, T. Maughan, S. Mukherjee, E.M. Hammond, Preclinical testing of an atr inhibitor demonstrates improved response to standard therapies for esophageal cancer, Radiother. Oncol. 121 (2016) 232–238. [33] A. Peasland, L.Z. Wang, E. 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Funding This work was supported by a grant from the National Institute of General Medical Sciences (GM130583to MGK). Declaration of Competing Interest None. Acknowledgments The authors thank the WSU Proteome Analysis Laboratory and Center for Genomics Research for the use of equipment to carry out this work. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.mrfmmm.2019. 111678. References [1] F. Valeriote, L. van Putten, Proliferation-dependent cytotoxicity of anticancer agents: a review, Cancer Res. 35 (1975) 2619–2630.
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ATR kinase inhibition sensitizes quiescent human cells to the lethal effects of cisplatin but increases mutagenesis
T
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Rebekah J. Hutcherson, Michael G. Kemp
Department of Pharmacology and Toxicology, Wright State University Boonshoft School of Medicine, 3640 Colonel Glenn Hwy, Dayton, OH, 45435, United States
A R T I C LE I N FO
A B S T R A C T
Keywords: Cisplatin Chemotherapy DNA damage response Quiescence Translesion synthesis Protein kinase signaling
The ATR protein kinase is known to protect cells from DNA damage induced during the replicative phase of the cell cycle. Small molecule ATR kinase inhibitors have therefore been developed to improve the effectiveness of DNA damage-based chemotherapy regimens aimed at killing rapidly proliferating tumor cells. However, whether ATR functions in a similar manner in non-replicating cells has not been examined and is important considering the fact that most cells in the body, including cancer stem cells in solid tumors, normally reside in either a quiescent or differentiated non-replicating state. Using cultured human cell lines maintained in a quiescent or slowly growing state in vitro, ATR was found to be activated following treatment with the common anti-cancer drug cisplatin in a manner dependent on the nucleotide excision repair (NER) system. Moreover, treatment with the ATR kinase inhibitors VE-821 and AZD6738 enhanced quiescent cell killing and apoptotic signaling induced by cisplatin. However, ATR kinase inhibition in quiescent cells treated with a low concentration of cisplatin also elevated the level of mutagenesis at the hypoxanthine phosphoribosyltransferase locus and resulted in increased levels of PCNA mono-ubiquitination. These results suggest that the excision gaps generated by NER may require a greater utilization of potentially mutagenic translesion synthesis polymerases in the absence of ATR kinase function. Thus, though ATR kinase inhibitors can aid in the killing of cisplatin-treated quiescent cells, such treatments may also result in a greater reliance on alternative mutagenic DNA polymerases to complete the repair of cisplatin-DNA adducts.
1. Introduction DNA damaging compounds are routinely used in the treatment of a variety of different tumor types. Rapidly proliferating cancer cells are generally thought to be at greater susceptibility to the lethal effects of DNA damaging drugs than normal cells and cells that are not actively progressing through mitotic cell cycle [1]. Furthermore, the ability of such compounds to induce cell death may be limited by the cellular DNA damage response (DDR), which is composed of diverse biochemical systems and signaling pathways that promote cell survival and recovery through DNA repair, cell cycle checkpoints, and other pathways [2–4]. The Ser/Thr protein kinase ATR (ataxia telangiectasia and rad3related) is a major regulator of the DDR, particularly in cells undergoing chromosomal DNA replication [5]. ATR has therefore recently emerged as a novel target for cancer chemotherapy regimens that are aimed at
improving the effectiveness of commonly used agents that generate DNA damage and replication stress [6–8]. Using diverse model organisms and systems ranging from yeast to frog egg extracts to cultured human cells, a plethora of studies have demonstrated that ATR limits replicating cells from the lethal effects of DNA damage by stabilizing stalled replication forks, inhibiting new replication origin firing, delaying the entry of cells into mitosis, enabling translesion synthesis, and promoting DNA repair and recombination [5]. Because nearly all of these events are specific to cells in S phase, our understanding of ATR function in the DDR is largely restricted to cells that are actively synthesizing DNA and progressing through the mitotic cell cycle. Given that most cells in the body are in a non-replicating quiescent or differentiated state, it is important to understand whether ATR can become activated in non-replicating cells and how ATR signaling impacts cellular responses to DNA damage that occur independent of canonical chromosomal DNA replication. For example, a recent study
Abbreviations: ATR, ataxia telangiectasia-mutated and rad3-related; NER, nucleotide excision repair; DDR, DNA damage response; TLS, translesion synthesis; HPRT, hypoxanthine phosphoribosyltransferase; SP, spironolactone; XPB, xeroderma pigmentosum group B; NA-AAF, N-acetyoxy-2-acetylaminofluorene; ssDNA, singlestranded DNA; RPA, replication protein A; PCNA, proliferating cell nuclear antigen ⁎ Corresponding author. E-mail address: [email protected] (M.G. Kemp). https://doi.org/10.1016/j.mrfmmm.2019.111678 Received 26 June 2019; Received in revised form 19 August 2019; Accepted 15 September 2019 Available online 17 September 2019 0027-5107/ © 2019 Elsevier B.V. All rights reserved.
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weeks as previously reported [11,12].
found that ATR inhibition can either promote cell death or survival in response to treatment with the bulky DNA adduct-inducing fluorene metabolite N-acetoxy-2-acetylaminofluorene (NA-AAF) depending on whether the cells are in a replicating/cycling or non-replicating/noncycling state [9,10], respectively. Whether these opposing functions for ATR are seen in response to commonly used anti-cancer drugs is not known. This lack of knowledge is a potential concern because ATR kinase inhibitors are entering clinical trials as adjuvants in cancer chemotherapy regimens. Thus, the toxicity and mutagenicity of chemotherapy drugs in non-replicating normal cells and tissues and quiescent cancer stem cells may be positively or negatively impacted by the addition of an ATR kinase inhibitor. Using cisplatin as a model anti-cancer drug, we investigated the impact of small molecule ATR inhibitor co-administration in non-replicating, quiescent human cells in vitro. We observed that ATR is capable of becoming activated in quiescent cells treated with cisplatin and that ATR kinase inhibition sensitizes quiescent cells to the lethal effects of cisplatin. Though this would be a favorable outcome in nonreplicating tumor cells in vivo, we also found that ATR inhibition increased the level of mutagenesis and resulted in increased monoubiquitination of PCNA, which may imply a greater reliance on the potentially mutagenic translesion synthesis (TLS) pathway to fill in the gaps generated by the NER machinery. Thus, ATR kinase inhibition may have both positive and negative effects on quiescent cell responses to DNA damaging compounds that are commonly used to treat human cancers.
2.3. Protein immunoblotting Cells were washed with cold PBS, scraped from the plate, and pelleted by gentle centrifugation. Cells were then lysed for 20 min on ice in 20 mM Tris−HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, and 1% Triton X-100. Following centrifugation in a microcentrifuge for 10–15 min at maximum speed, the soluble cell lysates were transferred to new tubes for immunoblot analysis. Chromatin-associated proteins were obtained from cells following two extractions with a modified cytoskeletal buffer (10 mM Tris−HCl (pH 7.4), 100 mM NaCl, 3 mM MgCl2, 1 mM EDTA, 1 mM Na3VO4, 10 mM NaF, and 0.1% Triton X100). Equal amounts of cell/chromatin extracts were separated by SDSPAGE, transferred to nitrocellulose, and then probed by immunoblotting using standard procedures. Primary antibodies included antibodies against ATR (sc-1887), actin (I-19), PCNA (sc-10) from Santa Cruz Biotechnology, phospho-p53 (Ser-15, 9284) from Cell Signaling Technology, phospho-ATR (Thr-1989, GTX128145) from GeneTex, and RPA70 (A300-421A) from Bethyl Laboratories. All primary antibodies were used at a 1:1000 or 1:2000 dilution in 1X TBST (50 mM Tris−HCl pH 7.4, 135 mM NaCl, and 0.1% Tween 20). Chemiluminescence was visualized with Clarity Western ECL substrate (Bio-Rad) or SuperSignal West Femto substrate (Thermo Scientific) using a Molecular Imager Chemi-Doc XRS + imaging system (Bio-Rad). Signals within the linear range of detection were quantified using Image Lab software (Bio-Rad) as previously described [9,10]. All experiments analyzing DNA damage response signaling were repeated at least two times, as indicated, and the average (and standard error) relative level of protein/phosphoprotein expression was determined and plotted.
2. Materials and methods 2.1. Cell culture Human immortalized HaCaT keratinocytes and U2OS osteosarcoma epithelial cells were cultured at 37 °C in a 5% CO2 humidified incubator in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% Fetal Clone III (Hyclone), 6 mM L-glutamine, 100 units/ml of penicillin, and 100 μg/ml of streptomycin. Cycling, replicating cells were typically plated at 30% confluence and treated the following day. Noncycling, quiescent cells were plated at 40–60% confluence, grown for 2 days in normal medium until the cells reached confluence, and then maintained for 2–3 days in DMEM containing 0.5% Fetal Clone III and penicillin/streptomycin before being subjected to experimental treatments. The ATR inhibitors VE-821 and AZD6738 (Selleckchem) were dissolved in DMSO and diluted 1000-fold to be used at the final concentrations of 10 μM and 3 μM, respectively. Spironolactone (SP; dissolved in DMSO) was used at a final concentration of 10 μM. Cisplatin (Sigma) was dissolved in PBS at a 3 mM stock concentration and then added to cell culture medium to obtain the indicated final concentrations. Cells were pre-treated for 2 h with SP and 30 min with the ATR inhibitors before addition of cisplatin to the culture medium. All compounds remained in the culture medium for all acute assays of survival, DNA repair, and kinase signaling.
2.4. DNA immunoblotting Bromodeoxyuridine (BrdU) was added to the culture medium at final concentration of 10 μg/ml for 15 min before harvesting the cells. Genomic DNA was purified using the GenElute Mammalian Genomic DNA Miniprep Kit (Sigma) and quantified using PicoGreen fluorescence (Invitrogen) on a Synergy H1 Hybrid Multi-Mode microplate reader (BioTek). DNA was immobilized on nitrocellulose, dried, and immunoblotted with antibodies against BrdU (Sigma B8434), cisplatinmodified DNA (CP9/19; Abcam ab103261), or single-stranded DNA (Millipore MAB3034) as previously described [10]. 3. Results 3.1. ATR kinase inhibition sensitizes replicating cells to cisplatin To confirm previous reports that ATR kinase inhibition sensitizes asynchronously growing human cells to the chemotherapeutic drug cisplatin, sub-confluent HaCaT keratinocytes and U2OS osteosarcoma cells in the logarithmic phase of growth were pre-treated with vehicle or the ATR inhibitor (ATRi) VE-821 for 30 min and then exposed to increasing concentrations of cisplatin. As shown in Fig. 1A, the inhibition of ATR potentiated the lethal effects of cisplatin treatment. These results demonstrate that ATR limits the effectiveness of cisplatin at killing proliferating HaCaT and U2OS cells. To compare the response of replicating cells to that of non-replicating, quiescent cells, sub-confluent proliferating HaCaT and U2OS cells and cells grown to confluence and maintained in medium with a low concentration of growth factors were pulsed with the nucleotide analog BrdU to measure the level of DNA synthesis by genomic DNA immunoblotting. As demonstrated in Fig. 1B, both the HaCaT and U2OS cells incorporated significantly reduced levels of BrdU when in the confluent, growth factor-deprived state. These results indicate that a quiescent or slowly growing cell state can be readily generated in vitro with these cell lines, and thus for simplicity we will refer here to both
2.2. Cell survival, cell death, and mutagenesis assays Crystal violet staining was used to monitor cell survival 3 days after cisplatin treatment as previously described [9,10]. Quantitation of trypan blue-positive cells was measured using a Countess II Automated Cell Counter (ThermoFisher). Measurements of sub-G1/G0 cells was carried out with an Accuri C6 flow cytometer following staining of ethanol-fixed cells with propidium iodide. For long-term assays of cell recovery and mutagenesis, drug-containing medium was replaced with fresh, low-serum medium after 2 days of treatment. For clonogenic survival assays, cells were then trypsinized 3 days later, re-plated at 100 cells per 100 mm plate, and then stained with crystal violet 10–14 days later to count surviving colonies. Cells with mutations at the HPRT locus were quantified by selection with 4 μM of 6-thioguanine for 2–3 2
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Fig. 1. Cisplatin induces cell death in both proliferating and quiescent human cells. Subconfluent, proliferating (A) HaCaT keratinocytes and (B) U2OS osteosarcoma cells were pre-treated with vehicle (DMSO) or the ATR inhibitor VE-821 (10 μM) for 30 min before treatment with the indicated concentrations of cisplatin. Surviving cells were stained with crystal violet 3 days later, and the stain was then solubilized and quantified using a spectrophotometer. Proliferating and quiescent (C) HaCaT and (D) U2OS cells were pulsed with BrdU for 15 min and then BrdU content in genomic DNA was measured via DNA immunoblotting to quantify the relative level of DNA synthesis. (E) HaCaT and (F) U2OS cells in either proliferating or quiescent state were exposed to increasing concentrations of cisplatin, and cell survival was measured 3 days later. The asterisks indicate significant differences (p < 0.05) in survival or DNA synthesis between treatment groups.
Immunoblotting of protein lysates from these cells revealed that both ATR itself [13,14] and the tumor suppressor protein p53 exhibited increased phosphorylation after cisplatin treatment (Fig. 2A), including at even lower concentrations of cisplatin (Supplementary Fig. 1). Pretreatment of the cells with two different small molecule inhibitors of ATR (VE-821 and AZD6738) largely blocked this response (Fig. 2A, B), which indicates that ATR is responsible for these phosphorylation events in quiescent cells. The canonical model for ATR kinase activation is that the stalling of DNA polymerases at DNA lesions results in the uncoupling of replicative helicase and polymerase activities [15], which subsequently leads to the generation of single-stranded DNA (ssDNA) that becomes coated with RPA to enable the recruitment and activation of ATR [16] in conjunction with the ATR-activating protein TopBP1 [5,17,18]. In
lines as being in a quiescent or non-replicating state. Non-replicating cells are generally considered to be much more resistant to DNA damaging agents than replicating cells [1]. Consistent with this notion, we observed that the non-replicating, quiescent HaCaT and U2OS cells required much higher concentrations of cisplatin to kill cells in comparison to their proliferating counterparts (Fig. 1C). 3.2. ATR kinase activity is stimulated in cisplatin-treated quiescent cells in a nucleotide excision repair-dependent manner We next examined whether the ATR kinase becomes activated in non-replicating cells treated with cisplatin. Quiescent HaCaT and U2OS cells were therefore exposed for 24 h to concentrations of cisplatin that had little effect on acute cell survival (60 μM and 30 μM, respectively). 3
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Fig. 2. Cisplatin induces ATR kinase activation in an NER-dependent manner in quiescent human cells. (A) Quiescent HaCaT and U2OS cells were treated for 24 h with the indicated inhibitors and with cisplatin (60 μM or 30 μM, for HaCaT and U2OS, respectively), and then immunoblotting was performed to detect the activation of ATR kinase signaling. The asterisk indicates a non-specific band on the p53 immunoblot in U2OS cells. (B) Quantification of 3 independent experiments performed as described in (A). (C) Quiescent HaCaT cells were treated with DMSO or 10 μM spironolactone (SP) for 2 h before treatment for 24 h with 60 μM cisplatin. Protein lysates were analyzed by immunoblotting, and the results from 3 independent experiments were quantified and graphed. The asterisks indicate a significant difference (p < 0.05) in ATR autophosphorylation following cisplatin treatment.
sensitized the quiescent cells to killing by cisplatin (Fig. 3A). Similar results were obtained with the ATR inhibitor AZD6738 (Fig. 3B), in cells treated with the related platinating drug carboplatin (Fig. 3C), with trypan blue staining to quantify cells with a loss of membrane integrity (Supplementary Fig. 2), and in telomerase-immortalized normal human fibroblasts (data not shown). These results demonstrate that ATR kinase signaling is important for quiescent cells to properly respond to cisplatin treatment. Because previous studies revealed that ATR promoted the death of quiescent cells treated with the fluorene metabolite NA-AAF [9,10], the current data further suggest that though cisplatin and NA-AAF both induce the formation of DNA adducts that can be targeted for removal by the NER machinery, the inhibition of ATR kinase signaling can have dramatically different effects on cell fate. Cisplatin-induced cell death may occur via apoptosis, and thus we next examined apoptotic signaling in quiescent HaCaT and U2OS cells treated with cisplatin and an ATR inhibitor. Using the cleavage of the caspase substrate PARP as a measure of apoptotic signaling, we observed that treatment with the ATR kinase inhibitor VE-821 resulted in more PARP cleavage in quiescent HaCaT cells exposed to a range of concentrations of cisplatin than in DMSO-treated cells (Fig. 4A,B). Similar results were obtained with the ATR inhibitor AZD6738 (Fig. 4C). Analysis of DNA degradation by flow cytometric analysis of propidium
contrast, several studies have reported that a major pathway for ATR kinase activation in UV-irradiated quiescent cells occurs via single stranded DNA (ssDNA) intermediates that are generated as a result of UV photoproduct removal by the NER machinery [19–22]. To examine whether NER is required for ATR kinase activation in quiescent cells treated with cisplatin, non-replicating HaCaT cells were pre-treated with vehicle or with spironolactone (SP), which inhibits NER by causing the rapid proteolytic degradation of the core NER protein XPB [11,23]. As shown in Fig. 2C and Supplementary Fig. 1, pre-treatment with spironolactone largely blocked the cisplatin-dependent increase in ATR and p53 phosphorylation. We conclude that ATR can become activated in non-replicating, quiescent cells in a manner dependent on the NER machinery. 3.3. ATR kinase inhibition sensitizes cisplatin-treated quiescent cells to undergo an apoptotic form of cell death To examine the possible functions of ATR kinase signaling in cisplatin-treated quiescent cells, HaCaT and U2OS cells maintained in a non-replicating state were treated with the ATR inhibitor VE-821 and then exposed to increasing concentrations of cisplatin. Cell survival was then quantified 3 days later. These experiments revealed that VE-821 4
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Fig. 3. ATR kinase inhibition sensitizes quiescent human cells to cisplatin and carboplatin. (A) Quiescent HaCaT and U2OS cells were treated with vehicle (DMSO) or 10 μM VE-821 and the indicated concentration of cisplatin for 3 days, and then surviving cells were stained and quantified. (B) Experiments were performed as in (A) except that the ATR inhibitor AZD6738 (3 μM) was used. (C) Cells were treated as in (A) except that carboplatin was used in place of cisplatin. The asterisks indicate significant differences (p < 0.05) in survival between treatment groups at the concentrations of cisplatin or carboplatin.
iodide-stained cells further confirmed these findings (Fig. 4D) Thus, we conclude that ATR kinase inhibition enhances the ability of cisplatin to induce cell death via apoptosis in non-replicating quiescent cells.
3.4. ATR kinase inhibition results in increased mutagenesis in quiescent cells treated with cisplatin Though DNA damage can exert toxic effects on acute cell viability, the damage can also have potentially mutagenic and long-lasting effects
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Fig. 4. ATR kinase inhibition potentiates apoptotic signaling in cisplatin-treated quiescent cells. (A) Quiescent HaCaT cells were treated with DMSO or VE-821 and the indicated concentration of cisplatin for 48 h, after which cell lysates were immunoblotted to detect apoptotic signaling. (B) Quantification of results from at least three independent experiments performed as in (A). (C) Quiescent HaCaT cells were treated DMSO, VE-821 (10 μM) or AZD6738 (3 μM) and 60 μM cisplatin for 48 h. Cell lysates were probed with the indicated antibodies to monitor apoptotic signaling. (D) Cells treated for 4 days were stained with propidium iodide and analyzed by flow cytometry. The asterisks indicate significant differences (p < 0.05) in the percentage of cleaved PARP and sub-G1/G0 cells between the ATRi- and DMSO-treatment groups.
Fig. 5. ATR kinase inhibition in quiescent cells does not impact proliferative potential but results in increased mutagenesis. (A) Schematic of experiments examining how quiescent cells recover from treatment with cisplatin and an ATR kinase inhibitor. Quiescent HaCaT cells treated with a low concentration (10 μM) of cisplatin in the absence or presence of the ATR inhibitor VE-821 were re-plated 5 days later at low density in the presence of growth factors to allow for measurements of (B) clonogenic survival and (C) mutagenesis at the HPRT locus. The asterisk indicates a significant difference (p < 0.05) in the number of HPRT mutants as determined in three separate experiments.
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on cell proliferation. Moreover, quiescent cells are frequently induced to proliferate upon cell stress or tissue damage, and thus we next designed experiments to determine how ATR kinase inhibition in the nonreplicating, quiescent state impacts the ability of cisplatin-treated cells to recover and proliferate. A schematic of this experimental design is provided in Fig. 5A and involved the use of a low concentration of cisplatin that does not lead to detectable decreases in acute cell survival (Figs. 1 and 3). Interestingly, using a classical clonogenic survival assay to monitor cell recovery, we observed similar numbers of surviving colonies in the absence and presence of the ATR kinase inhibitor VE821 (Fig. 5B). Thus, ATR inhibition does not appear to significantly impact the long-term proliferative capacity of quiescent cells treated with a low concentration of cisplatin. In addition to inducing cell death, DNA adducts may also lead to mutagenesis. We therefore used the classical HPRT mutagenesis assay to quantify the number of cells that develop mutations at the HPRT locus, which enables cells to become resistant to the toxic nucleotide analog 6-thioguanine. As shown in Fig. 5C, ATR kinase inhibition resulted in an approximately 2.5-fold increase in the number of cisplatintreated cells with mutations at the HPRT locus. We therefore conclude that although ATR kinase inhibition in quiescent cells treated with a low concentration of cisplatin does not lead to significant changes in colony forming ability, the loss of ATR kinase activity under these conditions promotes mutagenesis. 3.5. ATR kinase inhibition does not impact the removal of cisplatin-DNA adducts by NER Cisplatin treatment results in the formation of intra-strand cisplatinDNA adducts that are potentially lethal and mutagenic to cells if not removed by the NER machinery. To determine whether the enhancement in cisplatin toxicity and mutagenicity by ATR inhibition in quiescent cells is correlated with a defect in cisplatin-DNA adduct removal, non-replicating HaCaT cells were treated with cisplatin in the presence or absence of the ATR inhibitor VE-821 and then genomic DNA was analyzed for cisplatin-DNA adduct content by immunoblotting. As shown in Fig. 6, the inhibition of ATR kinase activity did not significantly affect the rate of cisplatin-DNA adduct formation or removal from quiescent cell genomic DNA. These results indicate that the cisplatin lethality and mutagenicity that is exacerbated by ATR kinase inhibition in quiescent cells is not due to a defect in the damage removal step of NER.
Fig. 7. ATR kinase inhibition leads to increased PCNA monoubiquitination in cisplatin-treated quiescent cells. (A) Quiescent HaCaT cells were treated for 24 h with DMSO or VE-821 and the indicated concentration of cisplatin. Cells were then fractionated to enrich for chromatin-associated proteins, which were analyzed by immunoblotting with the indicated antibodies. The asterisk in the RPA70 blot indicates a non-specific band, and the characteristic monoubiquitinated form of PCNA is labeled as PCNA-ub. (B) Quantitation of results from 3 independent experiments performed as in (A). (C) Cells were pre-treated with spironolactone (SP) before exposure to cisplatin for 18 h and fractionation to enrich for the monoubiquitinated form of PCNA. The asterisks on the graphs indicate significant differences in RPA and mono-ubiquitinated PCNA chromatin association between the treatment groups (p < 0.05).
3.6. Inhibition of ATR results in elevated PCNA monoubiquitination Though cisplatin-DNA adduct removal by NER was not impacted by ATR inhibition (Fig. 6), earlier data indicating that ATR kinase activation is NER-dependent (Fig. 2C) suggests that a step of NER that occurs after damage excision may be relevant to the function of ATR in promoting survival and limiting mutagenesis. To examine this further,
Fig. 6. The rate of removal of cisplatin-DNA intrastrand adducts is not affected by ATR kinase inhibition. (A) Quiescent HaCaT cells were treated 30 μM cisplatin and either DMSO or VE-821 and then genomic DNA immunoblotting was performed at the indicated time points to detect the presence of cisplatin-DNA adducts. Blots were re-probed to detect total DNA (B) Quantitation of results from 2 independent experiments performed as in (A).
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that is observed at the HPRT locus in cisplatin-treated quiescent cells with inactive ATR (Fig. 5) may be due DNA synthesis by mutagenic DNA polymerases at NER gaps. Though this process promotes cell survival, the gap filling process may therefore be more mutagenic. A schematic summarizing these results is provided in Fig. 8.
quiescent HaCaT cells pre-treated with either vehicle or the ATR inhibitor VE-821 were exposed to cisplatin and then fractionated to enrich for chromatin-associated proteins. As shown in Fig. 7A, the levels of RPA and PCNA, which are known to associate with the ssDNA gaps generated by NER [24–26], were elevated on chromatin in a cisplatin dose-dependent manner. RPA is the major ssDNA-binding protein in human cells, and PCNA is a replicative polymerase clamp protein that facilitates gap filling DNA synthesis during NER [24,25,27]. Interestingly, we also observed the presence of a characteristic monoubiquitinated form of PCNA that was readily apparent in cisplatintreated quiescent cells. This modification is frequently used as a biochemical indicator for activation of the translesion synthesis (TLS) pathway of DNA synthesis [28], in which specialized polymerases are recruited to DNA via ubiquitin-binding domains to carry out DNA synthesis across damaged nucleotides or at difficult to replicate genomic regions. Thus, the TLS pathway is likely required in quiescent cells to fill at least a sub-set of excision gaps generated by the NER machinery [29]. Interestingly, though treatment with the ATR kinase inhibitor did not impact total PCNA levels on chromatin and resulted in only moderately higher levels of chromatin-associated RPA after treatment with a low dose of cisplatin (Fig. 7B), the levels of monoubiquitinated PCNA were 2- to 3-fold higher in cells treated with the ATR inhibitor. Treatment of cells with spironolactone to inhibit NER largely blocked PCNA monoubiquitination (Fig. 7C). Preliminary studies with quiescent HaCaT cells exposed to UVB radiation has revealed a similar phenotype upon ATR kinase inhibition (Shaj and Kemp, unpublished). Together, these results indicate that ATR inhibition may result in a greater dependence on the TLS pathway to complete gap filling DNA synthesis. However, additional work is needed to determine how the loss of ATR signaling impacts recruitment of specific DNA polymerases to excision gaps. Nonetheless, the increased mutagenesis
4. Discussion Cisplatin remains a widely used chemotherapeutic agent for the treatment of a variety of solid tumors. However, because the various DNA damage responses regulated by ATR, including DNA repair, cell cycle arrest, and replication fork stabilization may limit the effectiveness of cisplatin, clinical trials are currently underway to test whether ATR inhibition can potentiate the cancer cell killing effects of cisplatin and other DNA damaging drugs that generate replication stress [6,8]. Preclinical models using cancer cell lines in vitro [30–34] and tumor xenografts in mice in vivo [35–38] support the concept that ATR kinase inhibition can potentiate the ability of cisplatin to slow or prevent tumor growth. A great deal of knowledge has been gained on ATR function in the response to replication stress induced by cisplatin and other DNA damaging agents. Though several studies have demonstrated that DNA damaging agents can activate ATR kinase signaling in quiescent or G1 phase cells [19–21,39–45], the actual function of this signaling in nonreplicating quiescent cells is largely unexplored. This is an important issue because most cells in the body, including cancer stem cells, frequently reside in a non-replicating quiescent state. Cancer treatments that eliminate such cells may therefore be most effective at preventing tumor recurrence [46], and thus ATR inhibitors could aid in this process. However, cancer drugs can also induce mutations that drive tumor recurrence and carcinogenesis, and thus care must be taken to design Fig. 8. Schematic summarizing the effect of ATR kinase inhibition in cisplatin-treated quiescent cells. Under normal conditions, cisplatin-DNA adducts (indicated by G < > G) are removed by the NER machinery, ATR becomes activated, and the excision gap can be filled in by DNA synthesis and ligation. However, when ATR kinase is inhibited, the gap filling process becomes more dependent on the translesion synthesis (TLS) pathway, which can result in mutagenesis or cell death.
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treatment regimens that maximize cancer cell killing while also minimizing toxicity and mutagenicity in normal cells. Here we used an in vitro model of cell quiescence to better understand ATR kinase function. We found that ATR becomes activated in non-replicating quiescent cells in vitro in a manner dependent on the NER (Fig. 2) and that inhibiting ATR under these circumstances improves the cell killing and pro-apoptotic effects of cisplatin in non-replicating cells (Figs. 3 and 4). These results indicate that similar to the situation in replicating cells, ATR protects non-replicating cells from the damage caused by cisplatin. Interestingly, these results are in striking contrast to similar experiments in which replicating and non-replicating cells were treated with the fluorene metabolite N-acetoxy-2-acetylaminofluorene (NA-AAF) [9,10]. These previous studies showed that though ATR kinase inhibition sensitized replicating cells to NA-AAF, ATR inhibitors protected non-replicating cells from the lethal and apoptotic effects of NA-AAF. Though both cisplatin and NA-AAF generate DNA adducts that can be repaired by the NER system, it is possible that the bulkier adducts induced by NA-AAF are a stronger block to transcription than those induced by cisplatin, and thus transcription stalling-dependent activation of ATR may play a more prominent role in promoting apoptosis in that context [41]. It will therefore be important to examine how ATR kinase inhibitors modify cellular responses to DNA damage caused by different carcinogenic and anti-cancer compounds in non-replicating cells to better understand the signals that activate ATR as well as the downstream processes that are regulated by ATR. Nonetheless, our results here along with previous results using ionizing radiation [45] indicate that combination radio- and chemotherapies involving cisplatin and an ATR inhibitor may be useful to kill both the replicating and non-replicating cells that comprise solid tumors in vivo. However, such treatments could exacerbate the potentially toxic effects of cisplatin and other DNA damaging agents on specific organ systems. Moreover, our evidence that ATR kinase inhibition increases the mutation frequency and PCNA monoubiquitination (a signal for recruitment of TLS polymerases) (Figs. 5 and 7) suggests that anti-cancer therapies utilizing ATR kinase inhibitors may increase the risk of mutagenesis and carcinogenesis within quiescent cancer stem cells or in other tissues of the body. Thus, the risk of secondary tumors arising as a result of cisplatin-based anti-cancer therapies may be elevated by treatment with an ATR kinase inhibitor and should be monitored accordingly.
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Funding This work was supported by a grant from the National Institute of General Medical Sciences (GM130583to MGK). Declaration of Competing Interest None. Acknowledgments The authors thank the WSU Proteome Analysis Laboratory and Center for Genomics Research for the use of equipment to carry out this work. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.mrfmmm.2019. 111678. References [1] F. Valeriote, L. van Putten, Proliferation-dependent cytotoxicity of anticancer agents: a review, Cancer Res. 35 (1975) 2619–2630.
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