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. Author manuscript; available in PMC: 2024 Oct 15.
Published in final edited form as: Toxicol Appl Pharmacol. 2023 Sep 20;477:116696. doi: 10.1016/j.taap.2023.116696

ATR activation by Cr-DNA damage is a major survival response establishing late S and G2 checkpoints after Cr(VI) exposure

Sophia Valiente 1, Casey Krawic 1, Anatoly Zhitkovich 1,*
PMCID: PMC10591798  NIHMSID: NIHMS1933466  PMID: 37734571

Abstract

Inhalation exposure to hexavalent chromium is known to cause lung cancer and other pulmonary toxicity. Cellular metabolism of chromium(VI) entering cells as chromate anion produces different amounts of reactive Cr(V) intermediates and finally yields Cr(III). Direct reduction of Cr(VI) by ascorbate (Asc), the dominant metabolic reaction in vivo but not in standard cell cultures, skips production of Cr(V) but still permits extensive formation of Cr-DNA damage. To understand the importance of different forms of biological injury in Cr(VI) toxicity, we examined activation of several protein-and DNA damage-sensitive stress responses in human lung cells under Asc-restored conditions. We found that Asc-restored cells suppressed upregulation of oxidant-sensitive stress systems by Cr(VI) but showed a strong activation of the apical DNA damage-responsive kinase ATR. ATR signaling was triggered in late S phase and persisted upon entry of cells into G2 phase. Inhibition of ATR prevented the establishment of late-S and G2 cell cycle checkpoints and did not lead to a compensatory activation of a related kinase ATM. Inactivation of ATR also strongly impaired viability of Cr(VI)-treated lung cells including stem-like cells and revealed a significant formation of toxic Cr-DNA damage at low Cr(VI) doses. Our findings identified a major Cr(VI) resistance mechanism involving sensing of Cr-DNA damage by ATR in late S phase and a subsequent establishment of protective cell cycle checkpoints.

Keywords: hexavalent chromium, chromate, ATR, DNA damage response, cell cycle

Introduction

Inhalation exposure to chromium(VI) has been firmly established to cause cancers and other respiratory tract toxicity in humans (DesMarais and Costa, 2019; Salnikow and Zhitkovich, 2008). Carcinogenic risks of Cr(VI) in humans were independent on tobacco smoking (Behrens et al., 2023) and showed a linear dose-dependence in large occupational cohorts with exposure to chromates (Gibb et al., 2000; Proctor et al., 2016), which is indicative of a directly genotoxic mode of action in cancer causation. Toxic and genotoxic effects of Cr(VI) are the result of biological damage caused by the products of its intracellular reduction which yields stable Cr(III). Reduction of Cr(VI) outside the cells is a detoxification process producing poorly membrane-permeable and unreactive Cr(III) complexes with proteins and small molecules. Cellular reduction of Cr(VI) is nonenzymatic and driven by direct electron transfer from small thiols and ascorbate (Asc) (Krawic and Zhitkovich, 2023). The reaction with Asc is the dominant metabolic process for Cr(VI) in lung cells in vivo (Standeven and Wetterhahn, 1992; Suzuki and Fukuda, 1990). Mechanistically, reduction of Cr(VI) by Asc involves the initial transfer of two electrons yielding Cr(IV) as the first intermediate whereas thiols are one-electron reducers generating more reactive Cr(V) intermediates. Cells in standard tissue cultures contain very small amounts of Asc and consequently, metabolize Cr(VI) via one-electron reduction reactions with glutathione and to a lesser extent, with cysteine (Krawic and Zhitkovich, 2023).

Cellular metabolism of Cr(VI) was found to cause oxidation of protein thiols (Myers and Myers, 2009; Myers, 2012) and the formation of DNA damage including several types of Cr-DNA adducts (Macfie et al., 2010; Voitkun et al., 1994; Quievryn et al., 2002). Cells contain different protein and DNA damage-sensing systems which activate specific stress signaling networks governing adaptative responses and cell fate decisions. The importance of specific forms of biological injury can be inferred from the strength of the activation of specific stress sensor systems and the magnitude of their effects on cell fitness and survival. Several studies in standard cell cultures found that Cr(VI) induced oxidant-sensitive responses involving cytosolic stress-activated protein kinases (Chen et al., 2009; Chuang et al., 2000; Wakeman et al., 2005) and AKT (Lal et al., 2009). However, the robustness of these responses in Asc-replete human lung epithelial cells, the main target of Cr(VI) adverse effects in inhalation exposures, remains unclear. Restoration of Asc levels in human cells resulted in the suppression of oxidative DNA damage by Cr(VI) (Reynolds et al., 2012) which was associated with the loss of activation of the apical DNA breaks- and oxidant-responsive kinase ATM (Luczak et al., 2016). The formation of DNA-ds breaks by Cr(VI) was replication-dependent (Ha et al., 2004; Reynolds et al., 2007a, 2009), leading to the subsequent accumulation of these toxic lesions in G2 phase (Reynolds et al., 2007a; Zecevic et al., 2009; Xie al., 2009). Treatments of different cells with Cr(VI) was also found to increase the percentage of cells in G2 phase (Chun et al., 2010; Reynolds at al., 2012), although a specific stress-signaling pathway that connects DNA damage, replication, and the activation of G2 checkpoint has not yet been established.

In this work using H460 human lung epithelial cells grown with different Asc levels, we investigated Cr(VI)-induced activation of several protein-and DNA damage-responsive pathways. We found that Asc-restored cells lost activation of oxidant-sensitive stress signaling pathways but showed a strongly elevated signaling by the apical DNA damage-responsive kinase ATR. H460 and HBEC3 normal lung stem-like cells were strongly dependent on ATR activity for survival after Cr(VI) injury. Cr-DNA damage triggered activation of ATR in late S phase and this genotoxic stress response remained elevated upon entry of cells into G2 phase. ATR-driven signaling was responsible for the establishment of late-S and G2 checkpoints after Cr(VI) treatments. Our findings identified ATR activation as a major protective response to Cr(VI), linking sensing of Cr-DNA damage in late S phase to the establishment of cell cycle checkpoints and cell survival.

Materials and Methods

Chemicals.

Dehydro-L-(+)-ascorbic acid (D8132), ascorbate phosphate (49752), sodium ascorbate (A7631), K2CrO4 (216615), RO-3306 (SML0569) and nocodazole (M1404) were purchased from Sigma-Aldrich. KU99533 (S1092), AZD6738 (S7693), AZ20 (S7050) and PD-0332991 (S1116) were from Selleckchem.

Cell culture and cell cycle synchronization.

H460 and HBEC3 cells were purchased from the American Type Culture Collection. Cells were grown in 20% O2/5% CO2 atmosphere using RPMI-1640 medium supplemented with 10% fetal bovine serum (H460) or a vendor-recommended synthetic media supplemented with growth factors (HBEC3). Stock solutions (10 mM) of potassium chromate [Cr(VI)] were prepared in sterile distilled water. Cr(VI) treatments alone and in combination with various inhibitors were performed in complete growth media. In cell cycle synchronization studies, H460 cells were treated for 16 h with 0.5 μM PD-0332991 for G1 arrest and for 16 h with 10 μM RO-3306 for G2 arrest. G1-arrested cells entered S-phase at 3 h after removal of PD-0332991.

Restoration of Asc levels in cells.

Elevation of cellular Asc was done either by short incubations of cells with dehydroascorbic acid (DHA) or by overnight growth of cells with ascorbate phosphate (pAsc). For Asc delivery into H460 using DHA, cells were incubated for 90 min in Krebs-HEPES buffer [30 mM HEPES (pH 7.5), 130 mM NaCl, 4 mM KH2PO4, 1 mM MgSO4, 1 mM CaCl2] containing 0.5 mM D-glucose and 5% fetal bovine serum. DHA incubations with HBEC3 cells were done in complete growth media for 2 h. After the initial dose dependence analyses, we selected 0.2 mM DHA for H460 and 0.4 mM DHA for HBEC3 cells to restore physiological concentrations of Asc prior to the addition of Cr(VI). In the experiments with pAsc, it was added directly to cells in their standard growth media. Concentrations of cellular Asc were determined by measurements as a specific fluorescent product formed after the reaction of cell extracts with 1,2-diamino-4,5-dimethoxybenzene (Reynolds and Zhitkovich, 2007b). Cell volumes were determined from forward scattering measurements using a flow cytometer (FACSCalibur, BD Biosciences).

Immunoblotting.

Cells were collected by scraping in cold PBS, centrifuged at 300xg for 10 min at 4°C, and washed in cold PBS. Cell pellets were resuspended a 2% SDS buffer [2% SDS, 50 mM Tris-HCl (pH 6.8), 10% glycerol] supplemented with Halt Protease and Phosphatase Inhibitors (Thermo Scientific, #78443) and heated at 99°C for 10 min. Samples were cooled to room temperature and lysates were cleared from the insoluble debris by centrifugation at 10,000xg for 10 min. Proteins (typically 20 μg) were separated on SDS-PAGE gels and then transferred onto ImmunoBlot polyvinylidene difluoride membranes (1620177, Bio-Rad) by either semidry transfer (<50 kDa proteins) or overnight wet transfer (larger proteins). Membranes were typically cut into a few strips to probe for proteins with different molecular weight. Primary antibodies were obtained from Cell Signaling [ATM (2873), phospho-ATR (30632), phospho-AKT (4060), AKT (2938), DNAPK (12311), histone H3 (9715), phospho-H3 (9701), HIF1α (14179), HIF2α (7096), L7A (2415) FOXM1 (5436), SOD1 (2770), fibrillarin (2639), RAD17 (8561), phospho-RAD17 (6981), CDK1 (9116), phospho-CDK1 (4539), CDC25B (9525), phospho-CHK1-Ser317 (2344), phospho-CHK1-Ser345 (2348), phospho-p53 (9284), p38 (9212), phospho-p38 (9211), RRM2 (65939), phospho-ERK1/2 (9101), phospho-JNK (4668), CHK2 (3440), phospho-CHK2 (2661) and CDC25A (3652)], Santa Cruz [p53 (sc-126), ATR (sc-1887) and cyclin A (sc-751)], Sigma-Aldrich [γ-tubulin (T6557)] and Abeam [phospho-ATM (ab81292) and phospho-DNAPK (abl8192)]. Primary antibodies were used at manufacturers’ recommended dilutions. Secondary antibodies were horseradish peroxidase-conjugated goat anti-mouse (12-349, Millipore) and goat anti-rabbit (7074, Cell Signaling). ECL Western Blotting Detection Reagent (RPN2232) was from GE Life Sciences.

S-phase checkpoint.

Engagement of S-phase checkpoint was assessed by measurements of the rates of DNA synthesis using the thymidine analogue EdU (Ortega-Atienza et al., 2016). After overnight growth in media supplemented with 200 μM pAsc, cells were treated with Cr(VI) for 3 h in early or late S phase and 10 μM EdU was added for the last 30 min of treatments. Cells were collected by trypsinization, fixed overnight in 80% ethanol at 4°C, washed with PBS and then incubated with 0.5% Triton X-100 in PBS for 30 min. After a wash with PBS, permeabilized cells were incubated in a Click-iT reaction mixture (Click-iT EdU-Alexa Fluor 488 Flow Cytometry Assay kit, Invitrogen, C10420) to fluorescently label DNA-incorporated EdU. Cells were washed PBS and DNA was stained with 0.5 μg/mL 7-aminoactinomycin D (BD Biosciences, 559925). After removal of free 7-aminoactinomycin D with PBS washes, cells were resuspended in PBS for flow cytometry. Fluorescence-Activated Cell Sorting (FACS) of DNA-and EdU-stained cells was performed using FACSCalibur (BD Biosciences). Data were analyzed using the Cell Quest Pro software.

Chromium assays.

Cr(VI) reduction was monitored by recording chromate absorbance at 372 nm every 20 s for 60 min at 37°C. Reduction reactions contained 20 mM MOPS, pH 7.4, 150 mM NaCl, 20 μM Cr(VI) and 200 μM ascorbate or ascorbate phosphate. Uptake of Cr(VI) were measured by graphite furnace atomic absorption spectroscopy (AAnalyst600 Atomic Absorption Spectrometer, PerkinElmer) using nitric acid extracts of cells (Krawic and Zhitkovich, 2018).

Cytotoxicity.

Cytotoxicity of Cr(VI) treatments was measured by the CellTiter-Glo luminescent assay (Promega, G7571). Cells were seeded (500-1000 per well) into optical bottom cell culture plates (ThermoFisher Scientific, 165305) and grown overnight before treatments. Cytotoxicity measurements were taken at specified times after removal of Cr(VI).

Colony formation.

460 cells were seeded onto 6-well plates (200 cells/well) and grown overnight in the presence of 200 μM pAsc. Next day, the medium was replaced and cells were treated with Cr(VI) for 72 h in media containing 200 μM pAsc in the absence or presence of 0.6 μM AZD6738. After Cr(VI) treatments, cells were grown in the regular medium+/−AZD6738 till visible colonies were formed. Colonies were stained with the Giemsa solution.

Statistics.

Differences between two groups were evaluated by two-tailed unpaired t-test. Multiple comparisons were performed using one-way ANOVA with the post-hoc Tukey’s test. Significance levels were set at p<0.05, p<0.01, and p<0.001.

Results and Discussion

We selected H460 human lung epithelial cells as our main cellular model as we have extensively used this line for characterization of toxic effects of Cr(VI) in the past and found it to closely recapitulate responses in primary human cells (Reynolds et al., 2007b; Reynolds et al., 2009; Luczak et al., 2019). Although H460 is a transformed cell line, it retained wt-p53 and showed normal activation of two main apical DNA damage-responsive kinases by their canonical inducers such as oxidative damage for ATM (Zhang et al., 2006; Rubis et al., 2019) and replication stress for ATR (Wong et al., 2012). Although in some biological contexts activated KRAS confers a variable degree of dependence on ATR and its downstream kinase CHK1 (Hattori et al., 2011; Igarashi et al., 2023), our initial viability tests showed that inhibition of ATR in KRAS-mutated H460 cells was not cytotoxic, indicating a normal state of ATR signaling. Inactivation of ATR also produced similar changes in DNA damage responses in H460 and primary human cells (Wong et al., 2012; Luczak et al., 2019). H460 cells can be readily synchronized in different cell cycle phases by inhibition of CDK kinases which are key physiological regulators of cell cycle progression (Duronio and Xiong, 2013).

Similar to other cultured cells, standard cultures of H460 are severely deficient in Asc containing only 5-15 μM of this Cr(VI) reducer corresponding to ~1% physiological levels of Asc in the lung and other tissues (Zhitkovich, 2020), which makes thiols the dominant reducers of Cr(VI) in these cells. Restoration of physiological levels of Asc in H460 and primary cells by a short preincubation with DHA prior to the addition of Cr(VI) resulted in the suppression of oxidative injury (Reynolds et al., 2012) and prevented the activation of the ATM-driven stress signaling pathway (Luczak et al., 2016). The loss of oxidative damage by Cr(VI) in Asc-restored cells was consistent with the absence of reactive Cr(V) intermediate during the kinetically dominant reduction of Cr(VI) by Asc in physiologically relevant mixtures with glutathione and cysteine in defined chemical reactions and in cells (DeLoughery et al., 2014). Asc is an important cofactor for multiple cellular proteins that utilize Fe(II) and 2-oxoglutarate for their enzymatic activity (Zhitkovich, 2020) and it is possible that growth of cultured cells with Asc further changes their responses to Cr(VI) through altered cell physiology.

Restoration of cellular Asc by supplementation with pAsc.

Prolonged incubations with DHA or Asc could not be used for a long-term maintenance of cellular Asc due to instability of both forms of vitamin C in media. Addition of ascorbate phosphate (pAsc), a stable and redox-inactive compound that becomes dephosphorylated inside the cells, represents an alternative method for restoration of cellular Asc. We tested the ability of pAsc to deliver Asc into H460 cells by the addition of this compound to the normal growth media for different periods of time. We found that pAsc produced a nearly linear increase in cellular Asc levels during 2 and 4 h incubations, with longer incubations for 6 and 24 h delivering only smaller further increases (Fig. 1A). The presence of pAsc in media for 48 and 72 h showed gradual decreases in cellular Asc in comparison to 24 h incubations (Fig. 1B), pointing to a limited ability of H460 cells to retain Asc as it has been observed in freshly isolated human lymphocytes and in cultured cells with restored Asc by other means (Zhitkovich, 2020). A comparison of Asc levels in cells immediately after 24-h long incubations with pAsc and following 3 h growth in the regular media confirmed a significant loss of cellular Asc irrespective of its initial levels (Fig. 1C). These results showed that in order to assess the role of Asc in Cr(VI) toxicity, cells either need to be continuously maintained in the presence of pAsc or treated with Cr(VI) for short periods of time immediately after pAsc incubations. In contrast to a fast reduction of Cr(VI) by Asc, pAsc was completely devoid of Cr(VI)-reducing activity (Fig. 1D), indicating that the presence of pAsc in media would not detoxify Cr(VI) outside the cells. Measurements of cellular Cr levels showed that growth of H460 cells in pAsc-supplemented media did not produce marked effects on uptake of Cr(VI) (Fig. 1E).

Figure 1. Effectiveness of pAsc in restoration of cellular Asc and lack of its direct reactivity with Cr(VI).

Figure 1.

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(A) Time-dependent accumulation of Asc in H460 cells incubated in the presence of 200 μM pAsc for 0-6 h (means±SD, n=3). (B) Cellular levels of Asc in H460 cells after prolonged incubations with 200 μM pAsc (means±SD, n=3). (C) Retention of Asc in H460 cells after removal of pAsc. Cells were grown with pAsc for 24 h and assayed for cellular Asc immediately (0 h) or after 3 h incubation in media without pAsc (means±SD, n=3, **-p<0.01, ***-p<0.001 relative to 0 h). (D) Cr(VI) is reduced by Asc but not by pAsc. The rates of reduction of Cr(VI) were measured at 37°C in MOPS buffer, pH 7.4. Each line includes average measurements from three replicates. SD values were smaller than 5% of the means and not shown for clarity. (E) Cr(VI) uptake by H460 cells grown for 24 h in the presence of different concentrations of pAsc. Cells were treated with 15 μM Cr(VI) for 3 h in the regular media without pAsc supplementation. Data are means±SD, n=3.

Robust activation of ATR but not other stress-sensitive pathways in Asc-restored cells by Cr(VI).

Protein and DNA damage activate largely nonoverlapping stress-sensitive signaling pathways that coordinate adaptation and cell fate-determining responses. High doses of Cr(VI) were previously found to cause oxidation of protein-SH groups (Myers and Myers, 2009; Myers, 2012), which was likely responsible for stimulation of stress-activated protein kinases (also known as mitogen-activated protein kinases) p38, ERK and JNK (Chen et al., 2009; Chuang et al., 2000). In order to assess a broad spectrum of stress responses to Cr(VI) in cells with different levels of Asc, we grew H460 cells for 24 h in the presence of 0, 50 and 200 μM pAsc and then treated them for 3 h with a cytotoxic dose of Cr(VI) (15 μM Cr, 34.6±3.5% viability at 48 h post-Cr, n=4). As observed earlier in standard cultures of other cells (Kaczmarek et al., 2007), Cr(VI) treatment of Asc-deficient H460 cells grown without pAsc supplementation produced a robust accumulation of hypoxia-inducible factors HIF1α and HIF2α (Fig. 2A), likely reflecting oxidative inactivation of functionally important SH-groups in the prolyl hydroxylase PHD2 controlling stability of HIF1α and HIF2α (Cyran and Zhitkovich, 2022; Briggs et al., 2016). Increasing cellular Asc to 0.36 mM by 50 μM pAsc supplementation was already sufficient to completely eliminate HIFα-stabilizing effects of Cr(VI). A modest increase in activating AKT phosphorylation by Cr(VI) was also detected only in Asc-deficient cells, potentially reflecting oxidative inactivation of its phosphatase PTEN (Zhang et al., 2020). Cells maintained in standard Asc-scarce culture had a strongly elevated background phosphorylation of p38 kinase, which was eliminated by elevation of cellular Asc to near physiological (50 μM pAsc addition) or physiological levels (200 μM pAsc addition) (Fig. 2A). Phosphorylation of p38 by Cr(VI) was evident in cells grown with 50 μM pAsc due to their lower background signal but this response became marginal in cells with physiological Asc (0.85 mM cellular Asc after growth in media with 200 μM pAsc). Two other stress-sensitive cytosolic kinases JNK and ERK1/2 did not show marked changes in their activating phosphorylation by Cr(VI) irrespective of Asc levels in cells. Metabolism of Cr(VI) in bacteria, which can be viewed as equivalent to one-electron reduction by thiols in Asc-depleted human cells, produces toxic superoxide radical and requires elevated SOD expression for survival (Branco et al., 2008). We found that protein levels of SOD1 in H460 cells were not affected by Asc supplementation (Fig. 2A), suggesting that the loss of the oxidant-sensitive responses in Asc-restored cells was probably unrelated to their enzymatic defenses against superoxide. Suppression of these responses in Asc-restored cells did not result from altered Cr(VI) accumulation as cellular Cr levels were not affected by the addition of pAsc (Fig. 1E).

Figure 2. Stress responses activated by Cr(VI) in cells with different levels of Asc.

Figure 2.

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In panels A and B, H460 cells were grown for 24 h in the presence of different concentrations of pAsc and then treated with 15 μM Cr(VI) for 3 h in the regular medium. (A) Activation of cytosolic stress-sensitive pathways (p-p38: T180/Y182-phosphorylated p38, p-JNK: T183/Y185-phosphorylated JNK, p-ERK1/2: T202/Y202-phosphorylated ERK1/2). (B) DNA damage-related responses. Bleo - cells were treated with the DNA oxidant bleomycin (5 μM, 3 h) and used as a positive control. (C) ATR-dependent and independent DNA damage responses in cells grown overnight in the presence of 200 μM pAsc, treated with Cr(VI) for 3 h in the regular medium and collected immediately. (D) ATR-dependence of DNA damage responses in cells treated as in panel C but collected for westerns 3 h after Cr removal.

Coordination of cellular responses to genotoxic stress is primarily driven by three apical kinases ATM, ATR and DNAPK (Blackford and Jackson, 2017; Ciccia and Elledge, 2010). ATM and DNAPK primarily respond to DNA damage in the form of DNA-ds breaks whereas ATR is activated by ssDNA which can arise after many forms of DNA damage. ATM can also be activated by its direct oxidation (Guo et al., 2010; Shiloh and Ziv, 2013) and in response to chromatin damage in the absence of DNA ds-breaks (Ortega-Atienza et al., 2016; Bakkenist and Kastan, 2003). As observed previously in Asc-deficient cells (Ha et al., 2003; Luczak et al., 2016), H460 cells in standard culture responded to Cr(VI) by activation of ATM, which was evident by strongly elevated phosphorylation levels of its two canonical targets: ATM-S1981 and CHK2-T68 (Fig. 2B). Elevation of Asc in cells by their growth in 50 and 200 μM pAsc-containing media led to a clear dose-dependent suppression of ATM-dependent phosphorylation of both targets. Activating autophosphorylation of DNAPK at S2056 was not affected by Cr(VI) at all cellular Asc levels, suggesting that ATM activation in Asc-deficient cells was likely caused by protein-damaging oxidative stress rather than oxidative DNA-ds breaks. In contrast to ATM, ATR-dependent phosphorylation assayed at three targets (ATR-T1989, CHK1-S317 and RAD17-S645) was strongly elevated by Cr(VI) in cells with different levels of cellular Asc (Fig. 2B). Cr(VI)-induced accumulation of the stress-sensitive transcription factor p53 was also largely insensitive to Asc levels whereas its Ser15 phosphorylation showed a modest decline in cells with high Asc, likely reflecting the loss of ATM contribution to this phosphorylation. Overall, our studies of stress-sensitive pathways with the high-dosage Cr(VI) showed the predominance of ATR-related genotoxic stress signaling in cells with restored Asc levels. Next, we investigated ATR and ATM-driven DNA damage responses in cells with restored Asc levels treated with 1.5-and 3-times lower Cr(VI) concentrations in the absence and presence of the ATR inhibitor AZD6738 (Fig. 2C). We found that these lower Cr(VI) doses also induced phosphorylation of p53 and CHK1 which were all completely dependent on ATR kinase activity. ATM activation measured by CHK2-T69 phosphorylation was barely changed in cells treated with Cr(VI) alone but showed a modest increase at 3 h post-Cr in ATR-inhibited cells. Formation of some more toxic DNA damage in ATR-inhibited cells could be responsible for this increase in CHK2 phosphorylation. Inhibition of ATR completely eliminated p53-S15 phosphorylation by Cr(VI) at 0 and 3 h post-exposure whereas p53 protein levels remained modestly elevated at the highest dose of Cr(VI), indicating a minor contribution of a non-ATR signaling to p53 stabilization via S15-independent phosphorylation. CHK1 and p53 phosphorylation remained strongly elevated at 3 h after Cr removal, demonstrating persistence of ATR-activating DNA damage (Fig. 2D). ATR inhibition did not induce a compensatory activation of ATM as evident by similarly low levels of CHK2 phosphorylation and the absence of p53-S15 phosphorylation in cells collected immediately and 3 h after Cr treatments (Fig. 2C,D).

Importance of ATR activation for survival of Cr(VI)-damaged cells.

Immunoblotting studies of Asc-restored cells treated with Cr(VI) showed a robust activation of ATR kinase (Fig. 2). To determine whether this response has protective or toxic consequences, we measured viability of pAsc-supplemented H460 cells exposed to Cr(VI) for 3 h and then grown for 48 h. We found that addition of ATR inhibitors (AZD6738 or AZ20) strongly diminished viability of cells treated with otherwise mildly-moderately cytotoxic doses of Cr(VI) (Fig. 3A). Consistent with its weak activation in pAsc-supplemented cells, inhibition of ATM kinase produced no significant effects on cytotoxicity of Cr(VI) at doses both below and above IC50 (Fig. 3B). Inhibition of ATR or ATM did not alter Cr accumulation in cells (Fig. 3C), excluding uptake differences as a confounding factor in cytotoxicity studies. A short preincubation with DHA represents a more rapid approach for elevation of cellular Asc levels to physiological levels (Fig. 3D). Viability measurements in Asc-restored cells by DHA preincubation also showed a strongly protective role of ATR after Cr(VI) damage but no significant impact of ATM kinase (Fig. 3E). Finally, we examined a long-term survival of pAsc-supplemented H460 cells following a continuous 72 h treatment with Cr(VI). We found that inhibition of ATR activity resulted in a dramatically lower colony formation by cells treated with low doses of Cr(VI) (Fig. 3F). Collectively, our cell viability studies showed that irrespective of the mode of Asc restoration in cells and duration of Cr(VI) treatments, ATR kinase played a major role in resistance of cells to Cr-DNA damage.

Figure 3. Importance of ATR but not ATM for viability of Asc-restored cells treated with Cr(VI).

Figure 3.

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H460 cells were treated with Cr(VI) for 3 h and viability measurements were taken 48 h later. Inhibitors were present during and after Cr(VI) treatments. (A) Viability of cells grown in the presence of 200 μM pAsc overnight and then treated with Cr(VI) in the regular medium in the absence or presence of ATR inhibitors (ATRi1 - 1 μM AZD6738, ATRi2 - 3 μM AZ20). Data are means±SD, n=4, ***-p<0.001 relative to Cr(VI) alone. (B) Viability of H460 cells treated as in panel A but in the absence or presence of 10 μM KU55933 (ATMi). Data are means±SD, n=4. (C) Cr(VI) uptake by H460 cells with inactivated ATM (10 μM KU55933, ATMi) and ATR (1 μM AZD6738, ATRi). Cells were treated as described in viability experiments (panels A and B). Data are means±SD, n=3. (D) Asc levels in H460 cells incubated with different concentrations of DHA. Data are means±SD, n=3. (E) Inhibition of ATR (1 μM AZD6738) but not ATM (10 μM KU55933) enhanced Cr(VI) toxicity in H460 cells preloaded with Asc using 200 μM DHA. Data are means±SD, n=4, ***-p<0.001 relative to Cr(VI) alone. (F) Colony formation by H460 cells grown overnight in the presence of 200 μM pAsc and then treated with Cr(VI) for 72 h in media supplemented with pAsc. ATR inhibitor AZD6738 (0.6 μM) was present during and after Cr treatments. Data are means±SD, n=3 (**-p<0.01, ***-p<0.001 relative to Cr alone).

ATR activation by Cr(VI) is triggered in late S phase and persists into G2 phase.

To determine cell cycle specificity of ATR activation by Cr(VI) under Asc-restored conditions, we took advantage of intact cell cycle checkpoints in H460 cells, which allows their synchronization by inhibition of CDK kinases. Inactivation of CDK4/6 kinases established a complete G1 arrest in H460 cells as evident by the absence of the S/G2 cyclin A and the S phase-inducible ribonucleotide reductase subunit RRM2 (Fig. 4A). Following removal of the CDK4/6 inhibitor, S phase was initiated at 3 h and completed at 9 h. After additional 3 h incubation (12 h after G1 release), cells entered mitosis as evidenced by a massive increase in histone H3-S10 phosphorylation. We found that Cr(VI) treatments of synchronized cell populations induced ATR-mediated phosphorylation of its target sites in CHK1 and p53 only in S phase (Fig. 4B), which in this experiment corresponded to the second half of S phase. To examine ATR activation by Cr(VI) during the entire S phase, we compared cells treated in early and late S phase. Analysis of three independent cell lysates for each condition found a consistently strong upregulation of ATR-mediated phosphorylation of its five targets and p53 stabilization in late S phase but only minor increases in early S phase for p53 and RAD17 with no appreciable changes for other phosphorylation sites (Fig. 4C,D). It appears that phosphorylation of p53 and its stabilization were induced slightly faster as they became modestly elevated already in early S whereas CHK1 phosphorylation was specific to late S phase. Striking differences in ATR activation by Cr(VI) in early and late S phase could have resulted from differences in DNA damage formation in early and late-replicating genomic regions or reflected a different sensitivity of cells to Cr-DNA damage. To investigate these possibilities, we treated early S-phase cells with Cr(VI) and analyzed DNA damage responses upon progression of these cells into late S and G2 phases (Fig. 4E). As observed above, Cr(VI) damage in early S phase (0 h post-Cr) did not trigger detectable CHK1 phosphorylation at its two ATR-targeted sites and again produced a modest increase in p53 phosphorylation (Fig. 4E, left panel). CHK1 and p53 phosphorylation by ATR became very abundant in cells analyzed at 3 and 6 h after Cr(VI) treatments, which corresponded to late-S and G2 phases, respectively. A large number of control cells collected at 6 h after mock treatments already entered mitosis as seen from a strongly elevated histone H3-S10 phosphorylation (Fig. 4E, right panel). In contrast, Cr(VI)-treated cells showed no evidence of mitotic cells at this time, indicating activation of strong checkpoint responses preventing progression of damaged cells into mitosis. Accumulation of G2 phase markers, such as inhibitory phosphorylation at CDK1-Y15 and accumulation of the mitosis-promoting transcriptional factor FOXM1 (Lindqvist et al., 2009), did not appear noticeably different between control and Cr(VI)-treated cells. Collectively, these studies showed that activation of ATR was initiated in late S phase irrespective of whether Cr-DNA damage was formed in early or late S phase, pointing to a heightened sensitivity of the ATR pathway during the completion of DNA replication cycle. ATR-driven stress signaling remained high after entry of Cr-damaged cells into G2 phase, which correlated with the block of progression of these cells into mitosis.

Figure 4. Late S phase activation of ATR by Cr(VI).

Figure 4.

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H460 cells were grown overnight in the presence of 200 μM pAsc and treated with Cr(VI) for 3 h in the absence of pAsc. (A) Progression of H460 cells through cell cycle following their release from G1 arrest (0.5 μM PD0332991 for 16 h). (B) Phosphorylation of ATR targets in different cell cycle phases by Cr(VI). H460 cells were synchronized in G1 by 16 h incubation with 0.5 μM PD0332991 and in G2 by 10 μM RO-3306 for 16 h. S-phase cells were treated with Cr(VI) at 6 h following their release from G1 block. (C) Phosphorylation of ATR targets in early S and late S phase assessed in three independent cell lysates in each group. Early S phase: Cr(VI) was added at 3 h after G1 release, late S phase: Cr(VI) was added at 6 h after G1 release. (D) Quantitation of early and late S-phase phosphorylation shown in panel C. Band intensities were measured by ImageJ and normalized to loading controls. Normalized values in untreated cells were subtracted from values in Cr-treated cells and the percentage of a specific phosphorylation in early and late S phase was calculated from the total phosphorylation in S phase (sum of early S+late S). Data are means±SD, n=3 (**-p<0.01, ***-p<0.001 relative to early S phase). (E) Immunoblots of cells treated with Cr(VI) in early S phase and collected at 0-6 h posttreatment.

Importance of ATR for S and G2 cell cycle checkpoints.

Functional consequences of activated ATR signaling in S and G2 phases could involve the establishment of cell cycle checkpoints in one or both of these phases. Engagement of S-phase checkpoint is manifested by the slowdown in the rate of DNA replication (Saldivar et al., 2017), which can be assessed by the DNA incorporation of the base analogue EdU. We found that Cr(VI) induced a robust checkpoint response in late S phase cells evidenced by their clearly decreased labeling with EdU (Fig. 5A). Consistent with the biochemical evidence for ATR activation in late S phase cells, ATR inactivation almost completely rescued inhibition of DNA synthesis in these cells by Cr(VI) indicating elimination of the checkpoint response. This conclusion was further supported by quantitative analyses of EdU labeling in late S phase cells (Fig. 5B). The inhibition of DNA synthesis in early S phase by Cr(VI) was less pronounced and it was only noticeable for cells with the highest EdU labeling (cells that were close to mid-S phase) with no apparent changes in cells with the intermediate labeling (early S-phase cells) (Fig. 5C). ATR inhibition slightly improved DNA synthesis by Cr(VI)-treated cells that were close to the mid-S phase. DNA synthesis in the early S-phase population of cells (intermediate EdU labeling) remained unaffected by Cr(VI) in cells with inactivated ATR. A complete lack of the DNA synthesis checkpoint in early S-phase and its modest engagement in cells close to the mid-S phase were consistent with the observed above kinetics of ATR activation by Cr(VI) (Fig. 4).

Figure 5. Role of ATR in S and G2 cell cycle checkpoints in Cr(VI)-treated cells.

Figure 5.

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H460 cells were grown overnight in the presence of 200 μM pAsc and treated with 5 μM Cr(VI) for 3 h in the absence of pAsc. Late S phase treatments: Cr(VI) addition at 6 h after G1 release, early S phase treatments: Cr(VI) addition at 3 h after G1 release. ATRi (2 μM AZD6738) was added together with Cr(VI) and was present during post-treatment incubations. (A) Representative FACS profiles demonstrating dependence of DNA synthesis inhibition in late S phase on ATR activity. Replication activity was determined by DNA incorporation of EdU which was added during the last 30 min of Cr(VI) incubations. (B) Relative replication rates measured from mean EdU signals in late S-phase cells (right peaks in FACS profiles) treated as in panel A. Data are means±SD, n=3 (untreated cells), n=5 (Cr-treated cells), ***-p<0.001, ns - not significant. (C) FACS profiles of EdU incorporation into early S phase cells treated with Cr(VI) in the absence and presence of ATRi. (D) Representative immunoblots of cells incubated for 3 and 6 h with the mitosis-trapping nocodazole added after Cr(VI) treatments. Cr(VI) was added to late S-phase cells (6 h after release from G1 arrest). (E) Quantitation of the mitotic marker phospho-histone H3 from the experiment shown in panel B. Data are means±SD, n=3, ***-p<0.001.

To monitor progression of Cr-damaged cells into M-phase, we employed a mitosis-trapping approach using the microtubule inhibitor nocodazole added after Cr(VI) treatments of late S-phase cells. ATR activity assayed by CHK1 and RAD17 phosphorylation was strongly elevated at 3 h after Cr(VI) removal (Fig. 5D). Phosphorylation of CHK1 moderately decreased at 6 h post-Cr but whereas levels of phospho-RAD17 remained similar. Progression of Cr-treated cells into mitosis was severely impaired as measured by the mitotic marker phospho-histone H3 (Fig. 5D,E). Strikingly, inhibition of ATR kinase almost completely rescued the inability of Cr-damaged cells to enter M-phase at 3 and 6 h post-Cr. At 3 h post-Cr, the absence of M-phase cells at least partially reflected activation of late S checkpoint (shown above by EdU labeling) and the resulting delayed entry into G2, with the latter being supported by a higher level of the S phase-acting phosphatase CDC25A in cells treated with Cr(VI) alone but not in the combination with the ATR inhibitor (Fig. 5D). The absence of the M-phase marker phospho-H3 at 6 h post-Cr can be attributed to the activity of G2 checkpoint as G2-specific biochemical changes such as the loss of CDC25A and the accumulation of CDC25B were fully established. A canonical mechanism for activation of G2 checkpoint by DNA damage such as DNA-ds breaks involves accumulation of the inhibitory Y15 phosphorylation at the M phase-promoting kinase CDK1 (Lindqvist et al., 2009; Shiloh and Ziv, 2013). DNA damage-activated ATR also typically arrest G2 cells by inactivating CDC25 phosphatases which reverse Y15 phosphorylation in CDK1 (Saldivar et al., 2017). We did not find marked changes in pY15-CDK1 levels in Cr-treated cells with normal or inactivated ATR (Fig. 5D). It is possible that the target of the ATR-driven G2 checkpoint after Cr(VI) is a subcellular localization of CDK1 (cytoplasmic retention) or some other CDK1-acting regulatory factor (Lindqvist et al., 2009). One such factor could be PLK1 kinase which was shown to be important for recovery from Cr(VI)-induced growth arrest in human fibroblasts maintained in standard tissue culture (Chun et al., 2010).

Protective role of ATR in HBEC3 human lung stem-like cells.

Cancers are believed to largely originate from malignant transformation of stem or progenitor cells (Blanpain, 2013). HBEC3 is a telomerase/CDK4-immortalized line of human bronchial epithelial cells with multipotent stem cell-like properties evidenced by their expression of epithelial markers of different lineage and the ability to differentiate into multiple types of lung cells (Delgado et al., 2011). Similar to H460 and other cultured cells, HBEC3 cells contain only very small amounts of Asc. Our attempts to restore physiological concentrations of Asc in HBEC3 cells using media supplementation with pAsc were unsuccessful whereas the addition of DHA effectively increased cellular Asc with a liner dose-dependence (Fig. 6A). The addition of 400 μM DHA delivered approximately 1 mM Asc into HBEC3 cells. In agreement with the results in H460 cells, we found that ATR inhibition in Asc-restored HBEC3 cells strongly enhanced cytotoxicity of Cr(VI), including the lowest dose that was noncytotoxic in cells with normal ATR activity (Fig. 6B). Immunoblotting for CHK1-S345 phosphorylation provided biochemical evidence for ATR activation by Cr(VI) in Asc-restored HBEC3 cells (Fig. 6C). A diminished viability of HBEC3 cells with inactivated ATR was not caused by Cr(VI) uptake differences, as cellular accumulation of Cr was not affected by the addition of ATR inhibitor (Fig. 6D). Thus, the DNA damage-responsive kinase ATR is also activated by Cr(VI) in normal lung stem-like cells and plays a major role in resistance of these cells to toxic effects of this carcinogen.

Figure 6. Importance of ATR for viability of Cr(VI)-treated HBEC3 cells.

Figure 6.

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(A) Concentrations of Asc in HBEC3 cells after incubation with DHA in growth media for 2 h. Data are means±SD, n=3. (B) Viability of HBEC3 cells treated with Cr(VI) for 3 h and assayed 72 h later. Cells were preincubated with 400 μM DHA for 2 h prior to the addition of Cr(VI). ATRi (1 μM AZD6738) was added together with Cr(VI) and present during post-Cr growth. Data are means±SD, n=4, ***-p<0.001. (C) Immunoblots of two sets of HBEC3 cells treated with Cr(VI) for 3 h as in panel B. (D) Uptake of Cr(VI) by HBEC3 cells treated with Cr(VI) for 3 h as panel B (means±SD, n=3).

Conclusions

Our studies here showed that Cr(VI) damage in H460 human lung cells with restored physiological levels of Asc did not trigger stress responses associated with protein damage (HIF1α/2α accumulation and four cytosolic stress-activated protein kinases as readouts). A shift of cellular Cr(VI) reduction to Cr(V)-skipping metabolism by Asc also suppressed activation of oxidant-sensitive and DNA damage-responsive kinase ATM. Cr(VI) metabolism in both Asc-deficient and Asc-restored cells strongly upregulated ATR-driven genotoxic stress signaling which was initiated in late S phase cells and persisted upon their progression into G2 phase. ATR activity was responsible for the establishment of late-S and G2 checkpoints which decreased the rate of DNA synthesis and blocked progression of Cr-damaged cells into mitosis, respectively. Survival of Cr(VI)-damaged cells with different modes of Asc restoration and duration of Cr(VI) treatments was strongly impaired by ATR inhibition, demonstrating a major role of this DNA damage-responsive kinase in protection against Cr(VI). These findings were further confirmed in human bronchial epithelial stem-like cells. In addition to the establishment of protective S and G2 checkpoints, other prosurvival activity of ATR in Cr-damaged cells may also include stimulation of DNA repair as suggested by the ATR-dependence of histone H2AX phosphorylation (DeLoughery et al., 2015).

Limitations of this study.

A main limitation of our work is the reliance on the use of transformed H460 cells for analysis of stress signaling responses. Although our previous studies and other reports found a normally functioning DNA damage response network in this line, the state of cytosolic stress sensors is less certain. H460 cells have mutated KEAP1 resulting in overexpression of the transcription factor NRF2 controlling the antioxidant response. Elevated enzymatic ROS defenses would not affect the formation and stability of Cr(V), the non-ROS oxidant produced during direct thiol-mediated reduction of Cr(VI), but they are expected to diminish the production of more potent oxidants such as Cr(V)-peroxo complexes (Krawic and Zhitkovich, 2023). On the other hand, high NRF2 and the resulting high levels of glutathione in H460 cells [2.7 mM as measured by a specific HPLC assay (Reynolds et al., 2012)] makes it more difficult to fully shift Cr(V)-generating prooxidant reduction of Cr(VI) by thiols to Cr(V)-skipping metabolism by Asc. Our use of pAsc also restored cellular Asc levels only to their low physiological range.

Supplementary Material

1

Highlights.

  • Cellular ascorbate suppressed activation of oxidant-sensitive responses by Cr(VI)

  • Cr(VI) strongly upregulated ATR signaling in ascorbate-restored human lung cells

  • ATR activity was induced in late S phase and remained elevated in G2 phase

  • ATR signaling was responsible for late S and G2 checkpoints by Cr(VI)

  • Survival of Cr(VI)-damaged lung cells was strongly dependent on ATR activity

Funding

This work was supported by grants ES031979, ES031002, ES028072 from the National Institute of Environmental Health Sciences.

Abbreviations:

Asc

ascorbate

DHA

dehydroascorbic acid

FACS

fluorescence-activated cell sorting

pAsc

ascorbate phosphate

Footnotes

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Credit Author Statement

Sophia Valiente: Investigation, Writing - Reviewing and Editing. Casey Krawic: Investigation, Writing - Reviewing and Editing. Anatoly Zhitkovich: Conceptualization, Supervision, Writing - Original draft preparation.

Declaration of interests

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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