The p53 gene is the most commonly mutated gene in solid tumors, but leveraging p53 status in therapy remains a challenge. Previously, we determined that p53 deficiency sensitizes head and neck cancer cells to AZD1775, a WEE1 kinase inhibitor, and translated our findings into a phase I clinical trial. Here, we investigate how p53 affects cellular responses to AZD1775 at the molecular level. We found that p53 modulates both replication stress and mitotic deregulation triggered by WEE1 inhibition. Without p53, slowing of replication forks due to replication stress is exacerbated. Abnormal, γH2AX-positive mitoses become more common and can proceed with damaged or underreplicated DNA. p53-deficient cells fail to properly recover from WEE1 inhibition and exhibit fewer 53BP1 nuclear bodies despite evidence of unresolved damage. A faulty G1–S checkpoint propagates this damage into the next division. Together, these deficiencies can intensify damages in each consecutive cell cycle in the drug.

Implications:

The data encourage the use of AZD1775 in combination with genotoxic modalities against p53-deficient head and neck squamous cell carcinoma.

This article is featured in Highlights of This Issue, p. 1013

Resistance to genotoxic therapy is the main reason that patients with head and neck squamous cell carcinoma (HNSCC) die of cancer, and evidence shows a strong association between loss of the p53 tumor suppressor and the emergence of resistance. HNSCCs have a very heterogeneous mutational landscape with few shared oncogenic mutations excepting the p53 gene (TP53), where mutations were noted in up to 72% of tumors (TCGA Research Network: http://cancergenome.nih.gov/; ref. 1). Despite the central role of TP53 in HNSCC carcinogenesis (2), to date no standard-of-care therapy leverages the tumor's p53 status, albeit preclinical work, and clinical trials are bringing this goal closer (3–5). We have previously found that inhibition of the cell-cycle kinase WEE1 with a small-molecule AZD1775 is significantly more cytotoxic to p53-mutated than to p53 WT HNSCC cell lines (6). Also, we recently completed a phase I trial of AZD1775 in combination with CDDP and docetaxel in HNSCC, which showed very promising results for patients with mutant or HPV-inactivated p53 (7). Our goal is to understand how p53 deficiency sensitizes HNSCC cells to AZD1775 as a single agent or in combination with genotoxic modalities.

WEE1 controls S phase and mitosis via inhibitory phosphorylation of cyclin-dependent kinases CDK2 and CDK1, respectively. Upon DNA damage or replication blockage, the ATM–CHK2 and/or ATR–CHK1 checkpoints block mitosis by acting on WEE1 and CDK1, thus allowing cells to complete DNA replication and repair. Inhibiting WEE1 can compromise the checkpoint, leading to forced mitosis and mitotic catastrophe (8–10). WEE1 inhibition also overactivates CDK2 during S phase, inducing replication stress through excessive initiation of replication and exhaustion of supplies of dNTPs, concomitant stalling of replication forks, and breakage of nascent DNA (11–13). Upon WEE1 inhibition, hyperactivation of CDK1/2 also suppresses RRM2 expression, exacerbating dNTP depletion (14), while precocious activation of CDK1 and PLK1 in S phase causes cleavage of stalled replication forks by the prematurely activated MUS81 endonuclease complex MUS81/SLX4 (15). The cytotoxic effect of the WEE1 inhibitor AZD1775 as a single agent is often attributed to induction of replication stress (16).

The prominence of mitotic and S-phase responses to AZD1775 and their relative contributions to the drug's cytotoxicity may differ depending on the cancer cells’ rewiring of the cell-cycle regulatory circuitry. Studies document different responses to AZD1775 in cell lines derived from sarcomas, carcinomas, leukemias, and other cancers (17–21). In some studies, S-phase arrest followed by the addition of AZD1775 promoted premature mitosis and cell death in the absence of p53 (8–10). However, in a study by Guertin and colleagues (22), induction of DNA damage in S phase, not premature mitosis, correlated with cytotoxicity of WEE1 inhibition in a panel of cell lines, and this effect was not dependent on the p53 status. Similarly, Van Linden and colleagues (18) noted no sensitization of AML lines to AZD1775 upon p53 inactivation.

Focusing on HNSCC cell models and isolating for p53-specific effects with an isogenic cell line pair, we previously reported p53-independent replication stress and p53-dependent unscheduled mitosis in an AZD1775-treated HNSCC cell line (23). Here, by following specific subpopulations of cells through more than one cell cycle, we reveal novel and confirm known p53-specific phenotypes in the response to WEE1 inhibition. Our results support the conclusion that an interplay of replication stress and G1–S and G2–M checkpoint failures can explain sensitivity of p53-deficient cells to AZD1775, and will help to optimize therapeutic window when targeting p53-mutated HNSCC.

Cell lines, vectors, and RNAi

Primary fibroblast cells (HFF4) were described previously (24). Head and neck cancer cell lines UM-SCC-74a was from Dr. Carey at University of Michigan (Ann Arbor, MI). Cells were used within one to 3 months after thawing and tested for Mycoplasma contamination prior to cryopreservation or upon thawing. We used a pBabeHygro retroviral vector expressing shRNA targeting p53 (ref. 25; a gift from Dr. Kemp) to generate a stable cell line with depleted p53 protein under hygromycin selection. siRNAs against p21 (CDKN1A) were from Qiagen (#SI00604898 and # SI00604905), and a nontargeting control siRNA (#D-001810-01-05) was from Dharmacon.

Drugs and chemicals

AZD1775 was provided by AstraZeneca through a collaborative agreement. CDDP (P4394) and Triapine (3-AP, SML0568) were purchased from Sigma-Aldrich. EmbryoMax Nucleosides (ES-008-D, EMD Millipore) were used at a final concentration of 1:25. 5-Iododeoxyuridine (IdU) and 5-chlorodeoxyuridine (CldU) were from Sigma-Aldrich and used at 50 μmol/L from stock solutions of 2.5 and 10 mmol/L in PBS, respectively.

Antibodies

Antibodies used were γ-H2AX (Ser139, JBW301 #05-636), p-HH3 (Ser10, 3H10, #05-806) from EMD Millipore; p-HH3 (Ser10, D2C8, #3377), p21 Waf1/Cip1 (12D1, #2947), cleaved PARP (D214, #9541), and β-Actin-HRP (13E5, #5125) from Cell Signaling Technology; P53BP1 (E-10, #sc-515841) and p53 (DO-1, #sc-126) from Santa Cruz; and nucleolin (#396400) from Life Technologies/Thermo Fisher. PE-conjugated anti-cleaved PARP antibody (Asp214 #51-9007684) was from BD Pharmingen. Antibody to IdU/BrdUrd (B44, #347580) was from BD Pharmingen and to CldU/BrdUrd [BU1/75 (ICR1), #OBT0030] from Bio-Rad/AbD Serotec.

Flow cytometry

Cells were fixed and processed for flow cytometry as described previously (24). Samples were run on FACSCanto II. FACS profiles were visualized using FACS Express software (DeNovo).

DNA fiber assays on sorted cells (Sorted Microfluidics-assisted Replication Track Analysis or SmaRTA)

Approximately 5 × 106 cells were fixed in 2% formaldehyde in PBS for 10 minutes at 37°C and stained with γH2AX and histone H3S10P antibodies as described for flow cytometry. DNA content was visualized by DAPI. Cells were sorted in PBS on Aria III sorter (BD Biosciences) to yield at least 50,000 cells per fraction. Sorted cells were pelleted after supplementing PBS with 0.3% BSA, resuspended in agarose plug buffer, embedded in agarose, and processed as described previously for the maRTA procedure (24, 26, 27).

Immunofluorescence and quantitative image-based cytometry (QIBC)

Cells were fixed and stained as described previously (24). QIBC was performed as described in ref. 28 with the following modifications: images were captured using TissueFAXS, an automated slide scanner (Zeiss AxioImager Z2 upright) microscope with a 20× objective. Automated image analysis for QIBC utilized TissueQuest software.

TP53 signaling pathway PCR array

The human p53 Signaling Pathway RT2 Profiler PCR array (Qiagen) was used as described by the manufacturer. UM-SCC-74a cells (p53wt vs. shp53) were treated with 1 μmol/L CDDP for 24 hours. Cells were harvested and processed according to the manufacturer's instructions.

Comet assays

DNA strand breaks were measured using a kit following the manufacturer's instructions (Trevigen). For each experimental condition, “tail moments” (defined as the product of tail length and the fraction of total DNA in the tail) were determined for at least 500 nuclei using ImageJ software (NIH) with the Open Comet plugin. At least 2 independent experiments were scored.

Cell growth and viability assays

Cell proliferation after drug washout was conducted as described previously (29). Alternatively, 103 cells were seeded into 96-well plates and were subjected to the same treatment for 24 hours. After drug removal, media were replenished and cells were allowed to grow for 96 hours before analysis with CellTiter-Glo (Promega), following the protocol outlined by the manufacturer.

Synergy analysis for drug combinations

A total of 103 cells were seeded into 96-well plates and treated with serial dilutions of the respective drugs for 96 hours. Cell viability was assessed with CellTiter-Glo (Promega), and the fraction affected was used to calculate the combination index (CI) and isobologram analyses according to the median-effect method of Chou and Talalay (30) using the CalcuSyn software (Biosoft).

Statistical analyses

Unpaired t tests were carried out in GraphPad Prism 7 software to analyze in vitro data. All data were expressed as mean ± SD (± SEM for large data sets), and P values were indicated. SmaRTA data were analyzed in Kolmogorov-Smirnov (K-S) tests using R studio software, and P values and, in some cases, D statistics are shown.

Cell-cycle progression and DNA-damage/replication stress response in WEE1 inhibitor-treated HNSCC cells

Phenotypes elicited by a therapeutically relevant dose of the WEE1 inhibitor AZD1775 were first demonstrated in the TP53 wild-type HNSCC line UM-SCC-74a. Pulse-chase labeling of cells with EdU and flow cytometry of EdU incorporation versus DNA content allows to see how EdU-positive cells progress through the cell cycle for up to two consecutive S phases (Fig. 1A; S1 and S2 in Fig. 1B). A majority of cells that were exposed to AZD1775 while in the S1 completed it and transited to G1 by 9.5 hours after EdU pulse, similar to untreated cells (Fig. 1B and C). The entry of the EdU+ population into the S2 and/or transit through it was delayed in WEE1-inhibited cells. In order to follow cell-cycle progression of AZD1775-treated cells past S1 more precisely, we consecutively labeled these cells with IdU and EdU as shown in Fig. 1C–E. The dual-labeled cells should be the ones that remained in the S1 phase for over 8 hours. We reasoned that these cells may be the most severely affected by AZD1775, and following their progression will reveal the strongest response to AZD1775. Figure 1D confirms the specificity of dual staining. Without the drug, a majority of IdU/EdU+ cells completed S1 and S2 phases, while with the drug these cells slowed a delay traversing through S2 (Fig. 1E). Together, the data suggest that response to WEE1 inhibition is heterogeneous and depends on the time in the inhibitor, indicating an accumulation of damage over more than one cell cycle.

Figure 1.

Response to WEE1 inhibition by AZD1775 develops over consecutive cell cycles in the HNSCC cell line UM-SCC-74a. A, Experimental design. Cells were pulse-labeled with EdU for 30 minutes, then grown for up to 36 hours with or without 300 nmol/L AZD1775. B, Flow-cytometric analyses. Left, representative density plots of cells stained for EdU incorporation and DNA content. Populations in the first S, G1, G2, and the second S phases since the EdU pulse are marked by arrows. Right, histograms of cell-cycle distribution of EdU-positive cells at the indicated times after the EdU pulse. C, Experimental design and a diagram of dual labeling: IdU is the first label and EdU is the second label. AZD1775 (400 nmol/L) was added after the first label where indicated. D, An example of immunofluorescent staining of dual-labeled, AZD1775-treated UM-SCC-74a cells harvested 15 hours after the second label. Scale bar, 40 μm. E, Histograms of cell-cycle distributions of dual-labeled cells at indicated times after the second label. F, Flow-cytometric analysis of cells treated as in A, harvested at indicated times of incubation with AZD1775, and immunostained for γH2AX expression, EdU incorporation, and DNA content. Top two rows are density plots of, respectively, γH2AX and EdU levels versus DNA content. The bottom row is histograms of cell-cycle distributions (by DNA content) of the following subpopulations: EdU-positive/γH2AX-negative (green), EdU-positive/γH2AX-positive (purple), γH2AX-superpositive (orange). G, Histograms of cell cycle (top) and EdU level (bottom) distributions of cells from the indicated subpopulations. Cells were incubated with AZD1775 for 24 hours and pulse-labeled for 30 minutes with EdU prior to harvest. H, A summary of findings presented in the figure. Red color intensity corresponds to the γH2AX level. Cells remain γH2AX-negative in early S phase, develop γH2AX signal as they progress through S, and at least some of them retain γH2AX staining in the next G1 and S. A subset of cells develops ultra-high level of γH2AX associated with suppressed DNA synthesis.

Figure 1.

Response to WEE1 inhibition by AZD1775 develops over consecutive cell cycles in the HNSCC cell line UM-SCC-74a. A, Experimental design. Cells were pulse-labeled with EdU for 30 minutes, then grown for up to 36 hours with or without 300 nmol/L AZD1775. B, Flow-cytometric analyses. Left, representative density plots of cells stained for EdU incorporation and DNA content. Populations in the first S, G1, G2, and the second S phases since the EdU pulse are marked by arrows. Right, histograms of cell-cycle distribution of EdU-positive cells at the indicated times after the EdU pulse. C, Experimental design and a diagram of dual labeling: IdU is the first label and EdU is the second label. AZD1775 (400 nmol/L) was added after the first label where indicated. D, An example of immunofluorescent staining of dual-labeled, AZD1775-treated UM-SCC-74a cells harvested 15 hours after the second label. Scale bar, 40 μm. E, Histograms of cell-cycle distributions of dual-labeled cells at indicated times after the second label. F, Flow-cytometric analysis of cells treated as in A, harvested at indicated times of incubation with AZD1775, and immunostained for γH2AX expression, EdU incorporation, and DNA content. Top two rows are density plots of, respectively, γH2AX and EdU levels versus DNA content. The bottom row is histograms of cell-cycle distributions (by DNA content) of the following subpopulations: EdU-positive/γH2AX-negative (green), EdU-positive/γH2AX-positive (purple), γH2AX-superpositive (orange). G, Histograms of cell cycle (top) and EdU level (bottom) distributions of cells from the indicated subpopulations. Cells were incubated with AZD1775 for 24 hours and pulse-labeled for 30 minutes with EdU prior to harvest. H, A summary of findings presented in the figure. Red color intensity corresponds to the γH2AX level. Cells remain γH2AX-negative in early S phase, develop γH2AX signal as they progress through S, and at least some of them retain γH2AX staining in the next G1 and S. A subset of cells develops ultra-high level of γH2AX associated with suppressed DNA synthesis.

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We next measured the γH2AX level as a function of DNA content and time in AZD1775. γH2AX accumulated over the course of 24 hours, with subsets of cells remaining γH2AX-negative (Fig. 1F, top). We also saw late-onset accumulation of extra-high level of γH2AX (γH2AX++) in a subset of mid S phase cells (2%–3% of all cells at 8.5 hours and 10%–14% of all cells at 24.5 hours, orange profiles, Fig. 1F, histograms). Appearance of γH2AX++ cells was not unique to the UM-SCC-74A line, as it was also observed in human primary fibroblasts and normal oral keratinocytes (Supplementary Fig. S1A) as well as in another TP53 wild-type HNSCC line, UM-SCC-81A (Supplementary Fig. S2E).

Tracking development of γH2AX by following EdU+ cells through the cycle (Fig. 1F, middle and histograms), we saw that early S phase cells were γH2AX and developed γH2AX signal as they reached mid S. A small subset of EdU+ cells was γH2AX as they reached G2 and traversed into G1. γH2AX level negatively correlated with the rate of progression through the S phase. Ninety-two percent of the γH2AX++ population at 8.5 hours in AZD1775 were EdU+ cells in their S1 phase in AZD1775. At 24.5 hours, in AZD1775 there were more γH2AX++ cells overall and only 65% of them were EdU+, suggesting an accumulation of cells in the γH2AX++ compartment over time (Fig. 1F and data not shown).

By labeling cells with EdU after 24 hours in AZD1775, we found that γH2AX++ cells incorporated about 10 times less EdU than γH2AX+ cells in S phase (Fig. 1G, histograms, compare orange and black profiles). γH2AX++ cells also expressed extra-high level of CHK1S345P (Supplementary Fig. S1B). Thus, these cells represent an S phase subpopulation with severely inhibited DNA synthesis and highly upregulated replication stress response. Findings of γH2AX induction are summarized in Fig. 1H.

Staining with an antibody against CDK1/2Y15P confirmed WEE1 inhibition (Supplementary Fig. S2A–S2D) and showed that at least in some cell lines γH2AX++ cells had a lower level of Y15 phosphorylation than γH2AX+ cells, potentially suggesting greater hyperactivity of CDK1/2 (Supplementary Fig. S2D and S2E). However, this lower staining for CDK1/2Y15P may be due to the fact that γH2AX++ cells are exclusively in the mid S phase, whereas γH2AX+ cells can be in the late S–G2, and CDK1/2Y15P staining is normally lower in mid S compared with G2 (Supplementary Fig. S2D).

Depletion of p53 exacerbates the response to AZD1775

We next depleted p53 in UM-SCC-74a background using shRNA (Fig. 2A). As expected, p53 depletion sensitized cells to AZD1775 (Fig. 2B). Depletion of p53 caused greater accumulation of γH2AX+ and γH2AX++ cells in AZD1775 (Fig. 2C and D). By immunofluorescence (IF) in situ (Fig. 2E), γH2AX++ expression level corresponded to extremely bright staining, either pan-nuclear or localized to numerous foci. Alkaline comet assays indicated that single- and double-strand DNA breaks (SSB and DSB) were elevated in AZD1775-treated p53 knockdown (kd) cells (Fig. 2F). Also, PARP1 cleavage (an apoptotic marker) was elevated in these cells (Fig. 2G and H).

Figure 2.

Depletion of p53 in UM-SCC-74a HNSCC cells modifies response to AZD1775. A, A Western blot verifies knockdown (kd) of the p53 protein and functional deficiency in p53 as demonstrated by the inability of cells to induce p21 after ionizing radiation. B, Fold change in proliferation relative to the untreated controls after treatment with 300 nmol/L AZD1775 for 16 hours in UM-SCC-74a and the primary fibroblast line HFF4. C, Flow-cytometric analysis of γH2AX response in mock and p53kd cells after 24 hours of 300 nmol/L AZD1775. D, Relative enrichment of γH2AX-positive cells in p53kd cells compared with the mock-depleted control (n = 3). E, Examples of immunofluorescent staining for γH2AX in AZD1775-treated p53kd cells. F, Alkaline comet assays in p53kd versus control cells treated 300 nmol/L AZD1775 for 16 hours (n = 3). G, Flow-cytometric analysis of cleaved PARP1. Fraction of cleaved PARP1 was assessed by gating cells relative to untreated controls and averaging the results (n = 9). H, A Western blot of cleaved PARP1 levels in cells treated with indicated doses of AZD1775 for 17 hours. I, Flow-cytometric analysis of EdU incorporation versus γH2AX expression. Black: all-negative cells; green: γH2AX-positive and superpositive (if any); blue: EdU-positive. Cells were labeled with EdU for 30 minutes and incubated for 24 hours with or without 300 nmol/L AZD1775. J, Histograms of cell-cycle distributions of EdU-positive cells pulse-labeled with EdU for 30 minutes and incubated with or without 300 nmol/L AZD1775 for the indicated times. EdU-positive cells that were γH2AX-negative (red) or γH2AX-positive (and superpositive, if any, blue) are plotted separately. K, Flow-cytometric analyses of EdU incorporation of cells incubated with AZD1775 for 24 hours with EdU labeling for 30 minutes prior to harvest. Low EdU-incorporating cells appearing upon AZD1775 treatment are marked by arrows.

Figure 2.

Depletion of p53 in UM-SCC-74a HNSCC cells modifies response to AZD1775. A, A Western blot verifies knockdown (kd) of the p53 protein and functional deficiency in p53 as demonstrated by the inability of cells to induce p21 after ionizing radiation. B, Fold change in proliferation relative to the untreated controls after treatment with 300 nmol/L AZD1775 for 16 hours in UM-SCC-74a and the primary fibroblast line HFF4. C, Flow-cytometric analysis of γH2AX response in mock and p53kd cells after 24 hours of 300 nmol/L AZD1775. D, Relative enrichment of γH2AX-positive cells in p53kd cells compared with the mock-depleted control (n = 3). E, Examples of immunofluorescent staining for γH2AX in AZD1775-treated p53kd cells. F, Alkaline comet assays in p53kd versus control cells treated 300 nmol/L AZD1775 for 16 hours (n = 3). G, Flow-cytometric analysis of cleaved PARP1. Fraction of cleaved PARP1 was assessed by gating cells relative to untreated controls and averaging the results (n = 9). H, A Western blot of cleaved PARP1 levels in cells treated with indicated doses of AZD1775 for 17 hours. I, Flow-cytometric analysis of EdU incorporation versus γH2AX expression. Black: all-negative cells; green: γH2AX-positive and superpositive (if any); blue: EdU-positive. Cells were labeled with EdU for 30 minutes and incubated for 24 hours with or without 300 nmol/L AZD1775. J, Histograms of cell-cycle distributions of EdU-positive cells pulse-labeled with EdU for 30 minutes and incubated with or without 300 nmol/L AZD1775 for the indicated times. EdU-positive cells that were γH2AX-negative (red) or γH2AX-positive (and superpositive, if any, blue) are plotted separately. K, Flow-cytometric analyses of EdU incorporation of cells incubated with AZD1775 for 24 hours with EdU labeling for 30 minutes prior to harvest. Low EdU-incorporating cells appearing upon AZD1775 treatment are marked by arrows.

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Flow-cytometric analyses of γH2AX development and cell-cycle progression showed that both p53kd cells and controls developed some γH2AX upon entry into their second S phase (S2) (Fig. 2I and J). Moderate expression of γH2AX during the S phase is not unusual for cancer cell lines. Most importantly, AZD1775 treatment delayed entry of control cells into S2, while p53kd cells entered S2 and expressed high level of γH2AX upon entry, as indicated by the appearance of an EdU-positive, γH2AX +, and ++ population at 24 hours in the drug (Fig. 2I).

Also of note, AZD1775 sped up the traversal of cells out of S1 and through G2-M compared with untreated controls (Fig. 2J, e.g., compare 4- and 8-hour time points with and without AZD1775). Compared with controls, p53kd cells had a higher G2 fraction both with and without AZD1775. The unrestrained entry into S2 observed in p53kd cells was also displayed by the HNSCC cell line PCI-15b harboring a high-risk TP53 mutation R273C (Supplementary Fig. S3A and S3B).

Pulse-labeling cells with EdU after a prolonged treatment with AZD1775 showed that both p53kd and control cells developed a mid-S phase population with severely depressed DNA synthesis (compare Fig. 2K with Fig. 1G; Supplementary Fig. S1A). Overall, the data suggest that p53 deficiency is associated with greater replication-associated damage upon AZD1775 treatment. At least part of this phenotype can be attributed to a failure of p53kd cells to activate the G1–S checkpoint and thus avoid an entry into their second S phase in the presence of the drug.

AZD1775 increases prevalence of mitosis in p53kd cells

WEE1 inhibition not only causes replication stress but also stimulates and in some cases advances mitosis (10, 31). To explore this facet of WEE1 inhibition in more detail, we stained cells for histone H3S10P modification as a marker of mitosis (Fig. 3). The correlation between the level of H3S10P staining and mitotic condensation and alignment of chromosomes was verified by IF (Fig. 3A). Of note, H3S10P level increased gradually in UM-SCC-74a cells and preceded visible chromosome condensation, suggesting that only the highest level of H3S10 phosphorylation identifies mitosis.

Figure 3.

Depletion of p53 in UM-SCC-74a cells increases prevalence of mitotic cells and leads to abnormal mitoses with high expression of γH2AX. A, Immunofluorescent staining for histone H3 phosphorylated on S10 (H3S10P) as a function of premitotic and mitotic stages. B, Flow-cytometric analysis of the H3S10P level versus DNA content. Cells were incubated with 500 nmol/L AZD1775 for 8 hours. C, Relative enrichment of high histone H3S10P (HH3+) cells in p53kd cells compared with the mock-depleted control (n = 3). D, Examples of immunofluorescent staining of AZD1775-treated mock and p53kd cells incubated with 300 nmol/L AZD1775 for 24 hours. Red: γH2AX; green: H3S10P. An arrow marks a cell staining positive for both markers. E, An example of an abnormal mitotic figure positive for both γH2AX and H3S10P in p53kd, AZD1775-treated cells. F, QIBC of mean fluorescent signals of γH2AX and H3S10P per nucleus in cells treated with 300 nmol/L AZD1775 for 24 hours. pB, n = 4,062; p53kd, n = 2,328. Dashed frames indicate expected positions of the γH2AX+/HH3+ subpopulation, and cells positive for both markers are marked by an arrow. G, Fractions of γH2AX-positive cells among the histone H3S10-positive, mitotic cells in p53kd cells and mock-depleted control. Treatment regimens were 300 nmol/L AZD1775/10 hours or 500 nmol/L AZD1775/8 hours. Cells were analyzed by flow cytometry (n = 5). H, A Western blot of siRNA-mediated depletion of p21 (CDKN1A) in control and p53kd cells. nc is nontargeting siRNA control. I, Levels of γH2AX+/++ and HH3+ cells relative to untreated control cells transfected with nontargeting siRNA, as measured by flow cytometry (n = 2). AZD1775 treatment was for 8 hours at 300 nmol/L. J, Fractions of γH2AX-positive cells among the histone H3S10-positive, mitotic cells in p53kd and control cells (n = 2).

Figure 3.

Depletion of p53 in UM-SCC-74a cells increases prevalence of mitotic cells and leads to abnormal mitoses with high expression of γH2AX. A, Immunofluorescent staining for histone H3 phosphorylated on S10 (H3S10P) as a function of premitotic and mitotic stages. B, Flow-cytometric analysis of the H3S10P level versus DNA content. Cells were incubated with 500 nmol/L AZD1775 for 8 hours. C, Relative enrichment of high histone H3S10P (HH3+) cells in p53kd cells compared with the mock-depleted control (n = 3). D, Examples of immunofluorescent staining of AZD1775-treated mock and p53kd cells incubated with 300 nmol/L AZD1775 for 24 hours. Red: γH2AX; green: H3S10P. An arrow marks a cell staining positive for both markers. E, An example of an abnormal mitotic figure positive for both γH2AX and H3S10P in p53kd, AZD1775-treated cells. F, QIBC of mean fluorescent signals of γH2AX and H3S10P per nucleus in cells treated with 300 nmol/L AZD1775 for 24 hours. pB, n = 4,062; p53kd, n = 2,328. Dashed frames indicate expected positions of the γH2AX+/HH3+ subpopulation, and cells positive for both markers are marked by an arrow. G, Fractions of γH2AX-positive cells among the histone H3S10-positive, mitotic cells in p53kd cells and mock-depleted control. Treatment regimens were 300 nmol/L AZD1775/10 hours or 500 nmol/L AZD1775/8 hours. Cells were analyzed by flow cytometry (n = 5). H, A Western blot of siRNA-mediated depletion of p21 (CDKN1A) in control and p53kd cells. nc is nontargeting siRNA control. I, Levels of γH2AX+/++ and HH3+ cells relative to untreated control cells transfected with nontargeting siRNA, as measured by flow cytometry (n = 2). AZD1775 treatment was for 8 hours at 300 nmol/L. J, Fractions of γH2AX-positive cells among the histone H3S10-positive, mitotic cells in p53kd and control cells (n = 2).

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High histone H3S10P-staining (HH3+) cells were more prevalent in p53kd cells compared with controls (Fig. 3B). AZD1775 markedly increased HH3+ abundance, particularly in p53-depleted cells (Fig. 3B and C). A minor fraction of HH3+ cells in the p53kd line also appeared to have < 4c DNA content (Fig. 3B). Notably, in the control, a vast majority of cells were either γH2AX+ or HH3+, whereas double-positive, HH3+/γH2AX+ cells were detectable in p53kd cells (Fig. 3D–G).

These differences were recapitulated in the p53-mutant PCI-15b cell line (Supplementary Fig. S3C, S3D, and S3E). In particular, basal and AZD1775-induced percentage of HH3+ cells was higher in these p53-mutant cells than in a p53 wild-type line, and a higher proportion of HH3+ cells was also γH2AX-positive. Furthermore, inactivation of p53 in UM-SCC-74a cells by expressing E6 protein of the HPV16 virus (32) recapitulated AZD1775-associated phenotypes displayed by p53kd cells, including suppression of growth, increased PARP1 cleavage, and elevated γH2AX and HH3+/γH2AX+ cells (Supplementary Fig. S4A–S4D). Thus, the data consistently show that induction of γH2AX expression and mitosis by WEE1 inhibition are more pronounced if p53 is altered, and second, that p53 deficiency correlates with the presence of mitotic γH2AX-positive cells. This feature may mark a specific vulnerability of p53-defective cells to WEE1 inhibition and/or may serve as a biomarker of WEE1i sensitivity.

The impact of p53 on the G2–M checkpoint can be conveyed through p21, a CDK inhibitor inducible by p53 (33, 34). Indeed when we depleted p21 in p53 wild-type cells, we observed an increase in γH2AX, HH3+, and HH3+/γH2AX+(++) cells, which negatively correlated with the p21 level (Figs. 3H–J). p21 depletion also exacerbated reduction of p21 levels observed in p53kd cells and increased γH2AX-positive mitoses, albeit less markedly than in p53 wild-type cells (Figs. 3H–J).

Recovery from AZD1775 triggers additional γH2AX induction in p53kd cells

We next asked whether recovery from AZD1775 was affected by p53 status. Cells were incubated with AZD1775 and then allowed to recover for up to 24 hours (Fig. 4A), and analyzed by IF in situ and QIBC. p53kd cells had a higher percentage of γH2AX-positive cells throughout the recovery (Fig. 4B). Over 30% of p53kd cells scored γH2AX-positive for 8 hours after removal of AZD1775 versus 5% of control cells. Moreover, a persistently higher fraction of γH2AX-positive p53kd cells displayed aberrant, severely misshapen or fragmented nuclei, consistent with failed segregation (Figs. 4C and D). However, this nuclear fragmentation was low overall, occurring in less than 10% of γH2AX-positive subpopulations throughout recovery.

Figure 4.

p53kd UM-SCC-74a cells recovering from AZD1775 retain more γH2AX but fail to form 53BP1 nuclear bodies. A, Experimental design. Cells were treated with 300 nmol/L AZD1775 for 24 hours, then allowed to recover for up to 24 hours. In some experiments, cells were also labeled with EdU for 30 minutes between 22.5 and 23 hours of the drug treatment. Cells were immunostained for γH2AX, H3S10P, and where indicated, EdU, and analyzed by QIBC. B, Relative enrichment of γH2AX-positive fraction in p53kd cells compared with controls (n = 2). Mean nuclear γH2AX signal was scored in at least 2,000 and up to 16,000 cells in each sample. Cells were considered γH2AX-positive if their mean γH2AX signal was >1/2 MAX of the population distribution, and the percentage of these cells was calculated for each sample, expressed as fold enrichment over the control at time point 0 hours recovery in each experiment, and averaged. Both γH2AX++ and γH2AX+ cells are included in this metric. C, Examples of immunofluorescent staining of p53kd, AZD1775-treated cells. The nucleus marked by an arrow displays aberrant morphology. D, Relative enrichment of cells with aberrant nuclear morphology among γH2AX-positive p53kd cells compared with controls measured in the same experiments as in B. Aberrant and normal γH2AX-positive nuclei were scored manually in digital images collected using a scanner microscope. A total of 120–600 nuclei were analyzed per sample. Percentage of aberrant among γH2AX-positive nuclei was calculated for each sample, expressed as fold enrichment over the control at time point 0 hours recovery in each experiment, and averaged. E, F, QIBC analysis of an experiment performed as in A. Mean nuclear EdU, γH2AX, and H3S10P signals were measured in 5,000–23,000 cells for each cell line/time point, and the data were subsetted based on EdU signal values. An EdU-low/negative subset has EdU values within the first quintile of a data set. EdU-high cells have EdU values >1/2 MAX of the data set. In E, γH2AX (top panels) and H3S10P (bottom panels) values in EdU-negative subsets of cells are plotted as a function of recovery time. In F, these same values are plotted for EdU-high subsets. Numbers above plots are percentages of γH2AX-positive and HH3+ cells at each time point. See Supplementary Fig. S4 for more on selection and validation of EdU, γH2AX+, and HH3+ value cutoffs. Diagrams on the right denote the inferred cell-cycle position of EdU-negative and EdU-high cells at the time of EdU pulse labeling. G, QIBC analysis of AZD1775-treated cells recovering from the drug for 24 hours. Mean nuclear 53BP1 and γH2AX signals were measured in approximately 5,700 each of control and p53kd cells. Elevated 53BP1 signal in controls compared with p53kd cells is marked with a bracket. H, Average population 53BP1 signal intensities were derived from QIBC measurements (n = 3). Cells were treated with AZD1775 as in A and released from the drug for 24 hours.

Figure 4.

p53kd UM-SCC-74a cells recovering from AZD1775 retain more γH2AX but fail to form 53BP1 nuclear bodies. A, Experimental design. Cells were treated with 300 nmol/L AZD1775 for 24 hours, then allowed to recover for up to 24 hours. In some experiments, cells were also labeled with EdU for 30 minutes between 22.5 and 23 hours of the drug treatment. Cells were immunostained for γH2AX, H3S10P, and where indicated, EdU, and analyzed by QIBC. B, Relative enrichment of γH2AX-positive fraction in p53kd cells compared with controls (n = 2). Mean nuclear γH2AX signal was scored in at least 2,000 and up to 16,000 cells in each sample. Cells were considered γH2AX-positive if their mean γH2AX signal was >1/2 MAX of the population distribution, and the percentage of these cells was calculated for each sample, expressed as fold enrichment over the control at time point 0 hours recovery in each experiment, and averaged. Both γH2AX++ and γH2AX+ cells are included in this metric. C, Examples of immunofluorescent staining of p53kd, AZD1775-treated cells. The nucleus marked by an arrow displays aberrant morphology. D, Relative enrichment of cells with aberrant nuclear morphology among γH2AX-positive p53kd cells compared with controls measured in the same experiments as in B. Aberrant and normal γH2AX-positive nuclei were scored manually in digital images collected using a scanner microscope. A total of 120–600 nuclei were analyzed per sample. Percentage of aberrant among γH2AX-positive nuclei was calculated for each sample, expressed as fold enrichment over the control at time point 0 hours recovery in each experiment, and averaged. E, F, QIBC analysis of an experiment performed as in A. Mean nuclear EdU, γH2AX, and H3S10P signals were measured in 5,000–23,000 cells for each cell line/time point, and the data were subsetted based on EdU signal values. An EdU-low/negative subset has EdU values within the first quintile of a data set. EdU-high cells have EdU values >1/2 MAX of the data set. In E, γH2AX (top panels) and H3S10P (bottom panels) values in EdU-negative subsets of cells are plotted as a function of recovery time. In F, these same values are plotted for EdU-high subsets. Numbers above plots are percentages of γH2AX-positive and HH3+ cells at each time point. See Supplementary Fig. S4 for more on selection and validation of EdU, γH2AX+, and HH3+ value cutoffs. Diagrams on the right denote the inferred cell-cycle position of EdU-negative and EdU-high cells at the time of EdU pulse labeling. G, QIBC analysis of AZD1775-treated cells recovering from the drug for 24 hours. Mean nuclear 53BP1 and γH2AX signals were measured in approximately 5,700 each of control and p53kd cells. Elevated 53BP1 signal in controls compared with p53kd cells is marked with a bracket. H, Average population 53BP1 signal intensities were derived from QIBC measurements (n = 3). Cells were treated with AZD1775 as in A and released from the drug for 24 hours.

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We also pulse-labeled cells with EdU 1 hour prior to release in order to mark replicating cells and follow them through recovery. γH2AX, EdU, and H3S10P signal intensities were manually collected in digital images for specific cell categories, i.e., mitotic, premitotic, or γH2AX++ (Supplementary Fig. S5A). This allowed us to define a range of signal intensities that was associated with a particular category. In parallel, we also collected γH2AX, H3S10P, and EdU signal intensities for thousands of individual nuclei using automated image acquisition and analysis.

We determined the distributions of γH2AX and histone H3S10P signal intensities in cells that displayed none/low (termed EdU) or high EdU incorporation when labeled just prior to release from AZD1775 (Fig. 4E). EdU cells correspond to mid S phase γH2AX++ cells as well as cells that were outside of the S phase at the moment of labeling (Figs. 1G and H; 2K). These cells most frequently displayed high levels of γH2AX (signal intensity >125) consistent with γH2AX++ status. The prevalence of these cells and their signal intensity declined over time in control but less so in p53kd cells. The majority of EdU cells were mitotic (H3S10P signal >75) in AZD1775 and before 4 hours of recovery, with subsequent wave of mitoses developing at 24 hours of recovery in both control and p53kd cells. Based on the overall EdU levels in the populations, between 12 and 24 hours of recovery most of control and p53kd cells have divided once (Supplementary Fig. S5A)

EdU-high signal is consistent with cells in the early to mid S phase at the point of release from AZD1775, and these cells were overwhelmingly γH2AX-negative in both cell lines (consistent with the data in Fig. 1F and G). Mitoses in these cells peaked at 8 to 12 hours of recovery, which was predictably later than mitoses in the EdU-negative cells. Interestingly, however, in the p53kd line, cells that were virtually γH2AX-negative in AZD1775 began to develop γH2AX signal at 24 hours after removal of the drug. This development followed completion of mitosis by several hours and thus is consistent with the entry of the EdU-high subpopulation into the next S phase. An independent experiment confirmed that EdU-high population of p53kd cells increased its γH2AX level at 24 hours compared with the 0 hours after the drug, unlike the mock-depleted controls (Supplementary Fig. S5B). Overall, the findings suggest that early S phase cells that display virtually no γH2AX response in AZD1775 nevertheless incur some kind of damage. In p53-deficient background, this damage persists for hours after AZD1775 removal and is revealed at the time point that is consistent with the entry into the next cell cycle.

We further addressed this by visualizing 53BP1 nuclear bodies (i.e., large, bright foci) in control and p53-deficient cells recovering from AZD1775. While in AZD1775, neither cell line had 53BP1 signal above background, consistent with other studies (35). At 24 hours of recovery, a subset of control cells clearly displayed elevated 53BP1 signal (Fig. 4G and H). Remarkably, p53kd cells displayed lower 53BP1 signal. In both cell lines, the majority of 53BP1-positive cells did not express γH2AX, arguing against 53BP1 colocalization with DSBs. Thus, both control and p53-deficient cells retain unresolved damage after mitosis; however, p53-deficient cells have an altered response to it.

γH2AX level in AZD1775-treated cells correlates with replication fork slowing

Our data suggest that γH2AX++ cells represent a qualitatively distinct subpopulation with severe replication stress (Figs. 1G, 2K and Supplementary Fig. S1). WEE1 inhibition by AZD1775 is known to cause replication fork slowing and stalling (12). We wanted to determine whether severe replication stress in the γH2AX++ population corresponded to the slowest forks. We were also interested to see if p53 status affected fork response to AZD1775, and if γH2AX++/HH3+ double-positive cells were quantitatively more affected than γH2AX++/HH3 cells, reasoning that premature mitosis of γH2AX++/HH3+ cells may be stimulated by their complete inhibition of replication, as suggested in ref. 15.

To answer these questions, we devised a combination of flow sorting and DNA fiber analysis (Supplementary Fig. S6). We sequentially labeled cells with CldU and IdU (with or without AZD1775) to mark ongoing replication forks, immunostained these cells with antibodies against γH2AX and H3S10P, and flow-sorted them. γH2AX, +, and ++ fractions were obtained for all cell lines. For p53kd cells, the γH2AX++ fraction was subdivided into HH3+ and HH3 populations (Fig. 5A).

Figure 5.

AZD1775-treated UM-SCC-74a cells expressing different levels of γH2AX experience different degrees of replication fork slowing. A, Experimental design. Cells incubated with 300 nmol/L AZD1775 and no-drug controls were labeled with consecutive 30 minutes pulses of CldU and IdU prior to harvest, then immunostained for γH2AX and H3S10P, and sorted into subpopulations prior to DNA isolation and stretching. The γH2AX++/HH3+ fraction was available only in p53kd cells. B, Examples of replication tracks of ongoing forks in p53kd cells. Extremely short CldU and IdU segments in forks in treated samples prompted us to measure total (C + I) lengths for each ongoing fork, as shown below the images. Two representative images for each condition were compiled to show more tracks. C, Ongoing fork track length distributions measured in the indicated γH2AX (white) and γH2AX/H3S10P (gray) subpopulations. Numbers of tracks analyzed for each sample are shown below the graph. D, Ongoing fork track length distributions derived from an independent experiment with p53kd cells. Designations as in C. E, A summary of differences in ongoing fork track lengths between γH2AX-negative and -positive subpopulations in p53kd cells and controls, derived from two independent experiments. The differences were expressed as a D statistic, i.e., the maximal difference between two cumulative distributions. D statistic values were calculated in K-S tests comparing each of the γH2AX-positive populations to the γH2AX-negative baseline for each cell line (AZD1775-treated). Differences with the D statistic of 0.12 and above were significant. F, Change in proliferation relative to the untreated controls after treatment with 300 nmol/L AZD1775 for 16 hours in the presence or absence of nucleosides in the media (n = 8).

Figure 5.

AZD1775-treated UM-SCC-74a cells expressing different levels of γH2AX experience different degrees of replication fork slowing. A, Experimental design. Cells incubated with 300 nmol/L AZD1775 and no-drug controls were labeled with consecutive 30 minutes pulses of CldU and IdU prior to harvest, then immunostained for γH2AX and H3S10P, and sorted into subpopulations prior to DNA isolation and stretching. The γH2AX++/HH3+ fraction was available only in p53kd cells. B, Examples of replication tracks of ongoing forks in p53kd cells. Extremely short CldU and IdU segments in forks in treated samples prompted us to measure total (C + I) lengths for each ongoing fork, as shown below the images. Two representative images for each condition were compiled to show more tracks. C, Ongoing fork track length distributions measured in the indicated γH2AX (white) and γH2AX/H3S10P (gray) subpopulations. Numbers of tracks analyzed for each sample are shown below the graph. D, Ongoing fork track length distributions derived from an independent experiment with p53kd cells. Designations as in C. E, A summary of differences in ongoing fork track lengths between γH2AX-negative and -positive subpopulations in p53kd cells and controls, derived from two independent experiments. The differences were expressed as a D statistic, i.e., the maximal difference between two cumulative distributions. D statistic values were calculated in K-S tests comparing each of the γH2AX-positive populations to the γH2AX-negative baseline for each cell line (AZD1775-treated). Differences with the D statistic of 0.12 and above were significant. F, Change in proliferation relative to the untreated controls after treatment with 300 nmol/L AZD1775 for 16 hours in the presence or absence of nucleosides in the media (n = 8).

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DNA was isolated from these fractions and subjected to our DNA fiber-stretching protocol, maRTA (Fig. 5B; ref. 27). AZD1775 treatment slowed fork progression in all cells, but, as we expected, fork progression rate negatively correlated with the γH2AX level (Figs. 5C and D), confirming that γH2AX++ cells experience the highest level of replication stress. In addition, the γH2AX+ subpopulation of p53kd cells had slower forks compared with γH2AX+ controls, and, overall, p53kd cells displayed a more pronounced reduction in fork progression in γH2AX-positive compared with γH2AX-negative subpopulations (Fig. 5E). Interestingly, fork progression in γH2AX++/HH3+ cells was not slower but in fact faster than in γH2AX++ cells (Fig. 5C and D). The data suggest that, while p53 status affects the severity of replication stress at the replication fork level, the p53kd-specific γH2AX++/HH3+ cells do not exhibit higher replication stress than γH2AX++/HH3 cells. Thus, the level of replication stress/suppression of replication alone cannot explain the prevalence of this double-positive subpopulation in p53kd cells.

The above data suggest that replication stress may be necessary, but it is not sufficient to induce a p53-specific survival defect. As an independent test, we asked if survival of p53kd cells was improved by supplementation of the media with extra nucleosides. Such supplementation is known to alleviate WEE1i-induced replication stress (12, 14). Indeed, the addition of nucleosides improved survival of the control but not p53kd UM-SCC-74a cells (Fig. 5F).

Synthetic lethality of AZD1775 in combination with CDDP or triapine

Only subsets of p53kd populations exhibited extreme dysfunction or cytotoxicity when exposed to the clinically safe doses of AZD1775 in the experiments above. In order to amplify cytotoxic outcome, and given that AZD1775 is being tested in combination with chemotherapy in clinical trials, we next explored the effects of combining AD1775 with other drugs.

Triapine (3-AP) is a potent inhibitor of ribonucleotide reductase (RNR) currently in clinical trials. A phase II study of 3-AP in metastatic HNSCC noted that the drug was well tolerated but was not effective as a single agent (36). We reasoned that 3-AP can exacerbate AZD1775-induced replication stress. On the other hand, CDDP, a standard-of-care drug for HNSCC, is a DNA cross-linker. In this case, the ability of AZD1775 to override the G2–M DNA-damage checkpoint induced by CDDP may enhance cell killing if the two drugs are combined.

The 3-AP and CDDP used at doses that had minimal to no effect on their own suppressed colony formation when combined with AZD1775 (Fig. 6A). Proliferation of AZD1775/3-AP–treated cells was more affected in the p53kd line than in the control (Fig. 6B), and the drugs showed weak (additive) interaction in p53kd cells and not in controls (isobologram analysis in Supplementary Table S1). The 3-AP dose that had no effect on γH2AX and H3S10P as a monotherapy dramatically enhanced γH2AX expression and virtually arrested S phase both in p53kd and control cells when combined with AZD1775 (Fig. 6C). While 3-AP/AZD1775 combination and AZD1775 alone increased the prevalence of cells with low to intermediate H3S10P levels (Fig. 6C, right), corresponding to premitotic cells (as before, see Fig. 4), only in p53kd cells did these drugs markedly induce premature mitosis (Fig. 6C, note the HH3+ high/<4c DNA content cell population marked by an arrow).

Figure 6.

Cotreatment with AZD1775 and triapine (3-AP) leads to mitosis with underreplicated DNA in a p53kd UM-SCC-74a cells. A, Colony formation of cells treated with the indicated drug combinations. B, Change in proliferation relative to the untreated controls after treatment with 300 nmol/L AZD1775 with or without 300 nmol/L 3-AP for 16 hours (n = 6). C, Flow-cytometric analyses of cells stained for γH2AX (top), H3S10P (bottom), and DNA content. Cells were treated with 300 nmol/L 3-AP and/or 300 nmol/L AZD1775 for 16 hours. HH3+ cells with less than 4N DNA content are marked with a black arrow. D, Alkaline comet assays performed on cells treated with AZD1775 and/or 300 nmol/L 3-AP for 16 hours (n = 3). E, As in D, except nucleosides were added to the media during treatments (n = 3). F, Change in proliferation relative to the untreated controls after treatment with 300 nmol/LAZD1775 and 300 nmol/L 3-AP for 16 hours in the presence or absence of nucleosides in culture media (n = 6).

Figure 6.

Cotreatment with AZD1775 and triapine (3-AP) leads to mitosis with underreplicated DNA in a p53kd UM-SCC-74a cells. A, Colony formation of cells treated with the indicated drug combinations. B, Change in proliferation relative to the untreated controls after treatment with 300 nmol/L AZD1775 with or without 300 nmol/L 3-AP for 16 hours (n = 6). C, Flow-cytometric analyses of cells stained for γH2AX (top), H3S10P (bottom), and DNA content. Cells were treated with 300 nmol/L 3-AP and/or 300 nmol/L AZD1775 for 16 hours. HH3+ cells with less than 4N DNA content are marked with a black arrow. D, Alkaline comet assays performed on cells treated with AZD1775 and/or 300 nmol/L 3-AP for 16 hours (n = 3). E, As in D, except nucleosides were added to the media during treatments (n = 3). F, Change in proliferation relative to the untreated controls after treatment with 300 nmol/LAZD1775 and 300 nmol/L 3-AP for 16 hours in the presence or absence of nucleosides in culture media (n = 6).

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The 3-AP/AZD1775 combination dramatically increased SSBs and DSBs in the DNA of p53kd cells (Fig. 6D). It is unlikely that these breaks were associated only with cells undergoing premature mitosis, as the latter comprised only a minor fraction (2.2% on average) of the entire population. The addition of extra nucleosides completely suppressed these breaks in the control and largely suppressed them in p53kd cells (Fig. 6E), as well as improved survival of control but not p53kd cells (Fig. 6F), in line with the findings in Fig. 5F.

We next looked into the effect of the CDDP/AZD1775 combination (Fig. 7). The combination had a more severe growth-suppressive effect and triggered a greater accumulation of SSBs and DSBs in p53kd cells compared with controls (Fig. 7A and B). Isobolograms of the combination showed a synergistic interaction for p53kd cells and no interaction for controls (Supplementary Table S2). AZD1775 and CDDP synergized in inducing γH2AX (Fig. 7C) and suppressed mitosis in the control but not in p53kd cells (HH3+cells in Fig. 7C). In order to visualize cell-cycle distribution in the CDDP/AZD1775-treated cells, the isogenic pair was treated with the drug combinations and then pulse-labeled with EdU prior to harvest (Fig. 7D). p53kd cells had a far greater accumulation of S and G2 cells in CDDP than the control, but in both cases addition of AZD1775 together with CDDP overrode it. However, unlike in the control, the combination triggered appearance of cells that had S phase-like DNA content but failed to incorporate EdU (about 10% of total population).

Figure 7.

Cotreatment with AZD1775 and CDDP (CDDP) leads to forced mitosis in p53kd UM-SCC-74a cells. A, Change in proliferation relative to the untreated controls after treatment with 300 nmol/L CDDP with or without 100 nmol/L AZD1775 for 16 hours (n = 5). B, Alkaline comet assays performed on cells treated with AZD1775 and/or 300 nmol/L CDDP for 16 hours (n = 3). C, Flow-cytometric analyses of cells stained for γH2AX (top), H3S10P (bottom), and DNA content. Cells were treated with 1 μmol/L CDDP and/or 300 nmol/L AZD1775 for 16 hours. HH3+ cells seen despite an ongoing DNA-damage response in p53kd cells are marked with a black arrow. D, Cell-cycle distribution of cells incubated with the indicated doses of CDDP and/or 300 nmol/L AZD1775 for 24 hours and pulse-labeled with EdU for 30 minutes prior to harvest. EdU-negative (orange) and -positive (blue) populations are graphed separately to distinguish G1, G2, and S phase cells. E, A heat map of the p53 Signaling Pathway RT2 Profiler PCR array analysis of cells treated with no drug or with 1 μmol/L CDDP for 24 hours. F, A quantitation of flow-cytometric analyses of cleaved PARP1 in untreated cells and cells treated with 300 nmol/L AZD1775 and 1 μmol/L CDDP combination for 16 hours (n = 3).

Figure 7.

Cotreatment with AZD1775 and CDDP (CDDP) leads to forced mitosis in p53kd UM-SCC-74a cells. A, Change in proliferation relative to the untreated controls after treatment with 300 nmol/L CDDP with or without 100 nmol/L AZD1775 for 16 hours (n = 5). B, Alkaline comet assays performed on cells treated with AZD1775 and/or 300 nmol/L CDDP for 16 hours (n = 3). C, Flow-cytometric analyses of cells stained for γH2AX (top), H3S10P (bottom), and DNA content. Cells were treated with 1 μmol/L CDDP and/or 300 nmol/L AZD1775 for 16 hours. HH3+ cells seen despite an ongoing DNA-damage response in p53kd cells are marked with a black arrow. D, Cell-cycle distribution of cells incubated with the indicated doses of CDDP and/or 300 nmol/L AZD1775 for 24 hours and pulse-labeled with EdU for 30 minutes prior to harvest. EdU-negative (orange) and -positive (blue) populations are graphed separately to distinguish G1, G2, and S phase cells. E, A heat map of the p53 Signaling Pathway RT2 Profiler PCR array analysis of cells treated with no drug or with 1 μmol/L CDDP for 24 hours. F, A quantitation of flow-cytometric analyses of cleaved PARP1 in untreated cells and cells treated with 300 nmol/L AZD1775 and 1 μmol/L CDDP combination for 16 hours (n = 3).

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A greater effect of CDDP on accumulation of cells in S and G2 in the p53kd line prompted us to measure contribution of p53 to the activation of the G2–M checkpoint regulators. We profiled expression of p53 pathway genes with or without CDDP treatment in the control and p53kd UM-SCC-74A cells (Fig. 7E). p53kd cells showed a markedly higher expression of S and G2–M checkpoint regulators (WEE1, CHK1/2, CDC25A), DNA-damage response (BRCA1/2), and replication (PCNA, MLH1, MSH2, CDK2, CCNE1) genes upon treatment with CDDP (right, Fig. 7E), which agrees with a more pronounced accumulation of these cells in S–G2 (Fig. 7D). As expected, p53kd cells failed to activate proapoptotic genes (BAX, FAS, CASP9, etc.; left, Fig. 7E). This confirms a greater G2–M engagement at the transcriptional level upon genotoxic stimuli specific to p53 depletion and is in agreement with the inability of p53kd cells to escape from the CDDP- and AZD1775-vulnerable S and G2 phases by arresting in the G1 phase (Fig. 7D). Consistent with this, only in p53kd cells did CDDP/AZD1775 combo clearly enhance apoptotic PARP1 cleavage (Fig. 7F).

Overall, the data indicate that combination treatments that enhance replication stress or inflict DNA damage in the setting of WEE1 inhibition can enhance cytotoxic outcome in p53-deficient cancer cells in positive correlation with, respectively, premature or forced mitosis.

Differential effects of p53 deficiency

WEE1 inhibition by AZD1775 elicits both p53-dependent and -independent phenotypes, some of which develop over a course of more than one cell cycle. For instance, S phase–associated DNA damage triggered by WEE1 inhibition (as revealed by γH2AX expression) is more prevalent in the second S phase of the time course of incubation with the clinically relevant dose of the drug. This suggests a carryover of a lesion or a particular cellular state from one cell cycle to the next. Interestingly, a recent study showed that WEE1 inhibition can result in an elevated CDK1 activity persisting throughout the G1 phase, which may affect DNA-repair and the G1–S transition (37). Consistent with their weakened G1–S checkpoint, p53kd (Fig. 2) and mutant (R273C; Supplementary Fig. S3) cells are more likely to enter their second S phase in AZD1775 than wild-type controls, and thus exhibit more damage as a population (Figs. 1 and 2; Supplementary Fig. S3).

In addition to this expected difference, we observed three more differential phenotypes. First, p53kd cells continue to experience effects of AZD1775 after its removal. This is most obvious in the subpopulation of cells that has not had a chance to develop γH2AX while in AZD1775 (Fig. 4F–H; Supplementary Fig. S5). This subpopulation undergoes mitosis and enters the next G1 on a similar schedule in the wild-type and p53-deficient lines; however, only in the latter it subsequently upregulates γH2AX. Interestingly, at this time, p53BP1 bodies are detected in control but less so in p53kd cells. It is possible that some type of DNA damage persists or becomes detectable after AZD1775 but it is not properly responded to by p53-deficient cells. Alternatively, the type of damage that persists in p53kd cells is invisible to 53BP1. While the latter cannot be ruled out, comet assays suggest that both in control and p53kd cells the carryover damage may be derived from SSBs and gaps (visible to 53BP1); and these are in fact more prevalent in AZD1775-treated p53kd cells (Fig. 2). p53BP1 bodies have been implicated as one of the contributors to the G1–S checkpoint activation by p53 WT cells (38). However, to our knowledge, virtually no evidence (with one exception; ref. 39) thus far points at p53 as a factor contributing to the formation of the 53BP1 bodies despite the original finding of association between 53BP1 and p53 (40). Thus, our finding may suggest a previously undetected interplay between p53 and 53BP1 in regulating the G1–S checkpoint in the aftermath of WEE1 inhibition in HNSCC.

WEE1 inhibition is known to slow replication fork progression (12). By performing DNA fiber analyses on subpopulations of cells, we found another differential phenotype of AZD1775-treated p53kd cells: while exhibiting the same γH2AX response as controls, they had a more severe replication fork slowing compared with their respective γH2AX-negative baseline (Fig. 5). This novel observation can imply that on a cell-by-cell basis, γH2AX response to replication stress is actually dampened by the knockdown of p53, and/or that p53 facilitates fork progression under stress. A stimulatory role of p53 in stressed fork progression was found by some studies (41, 42) but not others (43). Resistance to AZD1775-induced replication stress at a replication fork level likely involves a RAD18–TLS polymerase kappa-dependent tolerance pathway and a RAD51-dependent fork protection pathway (13). We propose that p53 may regulate a choice between these pathways both directly (42, 43) and in a transcription-mediated manner (44).

Lastly, we observed that p53kd and p53 R273C-mutant cells were more likely than the wild-type to undergo mitosis in AZD1775 despite an ongoing DNA-damage response (Fig. 3; Supplementary Fig. S3). This forced mitosis manifested as increased prevalence of HH3+ cells, and moreover, it was associated with the appearance of a unique subset of γH2AX++/HH3+ cells, as well as a higher percentage of micronuclei and abnormal (lobed, broken) nuclei, which are typically associated with mitotic catastrophe. We previously reported (23) that a small fraction of γH2AX++/HH3+ p53-deficient cells displays less than 4c DNA content, i.e., enters mitosis with underreplicated DNA, and in this study, we demonstrate that this fraction of cells in mitotic catastrophe can be increased by interfering with DNA replication during AZD1775 treatment (Fig. 6). Similar findings have been recorded by other labs (10, 17, 20, 35), consistent with the notion that p53 dysfunction weakens the G2–M checkpoint. This impact of p53 on the G2–M checkpoint can be conveyed through p21, whose mitotic role has come into focus recently (33, 34, 45). Our data (Fig. 3) and (10), indeed, suggest that p21 contributes to preventing the G2–M checkpoint override by AZD1775. However, the p53–p21 axis is unlikely the only route by which p53 may regulate the onset of mitosis. p53 may also inhibit transcription of Aurora A, a kinase required for mitosis (46), or of FBW7, a negative regulator of Aurora B, another mitotic kinase (47).

p53-dependent mitotic deregulation has been reported predominantly in cells derived from carcinomas of the breast, colon, and head and neck and is associated with p53 knockdown, TP53-null mutations, and, interestingly, only a subset of cancer-associated TP53 missense mutations (10, 20, 22, 35, 48). This exposes a need for a better understanding of molecular activities of different p53 mutants in an otherwise isogenic context of specific cancer types as they respond to cell-cycle deregulation.

Interaction between replication stress and mitotic deregulation, and the insights into combination treatments with AZD1775

Our results are consistent with the notion that replication stress and mitotic deregulation are independent variables in the response to AZD1775, and that replication stress is not sufficient to confer p53 sensitivity to AZD1775 in HNSCC. First, HH3-positive and HH3-negative p53kd cells exhibited similar levels of replication stress as measured by their γH2AX expression and the extent of their replication fork slowing (Fig. 5), which suggests that a property other than the severity of replication stress determined whether these cells underwent mitosis. Second, relieving replication stress by supplementation of extra nucleosides in the media did not improve survival of p53kd cells (in contrast to controls), while reducing such telltale symptom of replication stress as SSBs in DNA (Figs. 5 and 6). The ability of nucleosides to improve replication fork progression and suppress phenotypes of AZD1775-induced replication stress is well documented (12, 35).

At the same time, it is clear that replication stress can be manipulated to work together with mitotic deregulation in p53kd cells in order to boost cytotoxicity. For example, combining AZD1775 with a relatively novel, therapeutically relevant replication blocker triapine (3-AP), increased premature mitosis of underreplicated DNA and additively suppressed survival of p53kd HNSCC cells (Fig. 6).

A standard-of-care chemotherapy for HNSCC, CDDP represents another approach to boosting cell killing by AZD1775. CDDP does not prevent complete replication of the genome but slows it through engagement of the S phase checkpoint. CDDP induces cell death or senescence, and successful repair of CDDP damage heavily relies on homologous recombination in the S and G2 phases of the cell cycle and thus depends on a functional G2–M checkpoint (49). In agreement with this, we observed that when treated with CDDP, p53kd cells (incapable of G1 arrest) showed a greater induction of S and G2–M damage checkpoints than controls (Fig. 7). AZD1775 overrode this response, with concomitant induction of mitosis and PARP1 cleavage indicative of apoptosis. Together, these molecular phenotypes explain synergistic interaction between CDDP and AZD1775 that we observed in p53kd HNSCC.

Our results are similar, though not identical to the findings of Osman and colleagues (48). For example, p53-defective cells were not hypersensitive to AZD1775 alone in the PCI-13 background used by the authors, while we detected sensitization of the UM-SCC74a cells by p53 depletion or inactivation. The variability is likely an example of cell line heterogeneity; nevertheless, taken together the results point to a promising therapeutic potential of combining the standard-of-care CDDP with WEE1 inhibition in p53-defective HNSCC.

Intrapopulation heterogeneity of responses to AZD1775: A p53-independent phenotype

We observed significant heterogeneity in the γH2AX response of cells to AZD1775 and demonstrated that it is associated with major functional differences among the cells. In particular, we detected a pronounced negative correlation between cellular γH2AX levels and rates of replication fork progression, which brings up the question why subsets of cells experience higher or lower replication stress in AZD1775. Because γH2AX-negative cells in AZD1775 typically belonged to early S phase, we hypothesize that their γH2AX-negative state is linked to an intrinsically low level of CDK1 at this point of the cycle. Alternatively, a critical stress-signaling lesion may be slow to accumulate in these cells.

Extreme overactivation of CDK1 (and possibly CDK2) may also help explain why a subset of γH2AX-positive cells (γH2AX++) in mid to late S phase go on to manifest extremely high replication stress (Figs. 1, 2, and 5; Supplementary Figs. S1 and S2). Indeed, a γH2AX-positive subpopulation with similar properties was previously observed upon overexpression of CDK1/2-activating phosphatase, CDC25A (50). If so, it will be of interest to understand how and why some cells within a population suffer a more severe overactivation of CDK1/2 than others, and find ways to utilize this knowledge in cancer therapy.

In summary, our high-resolution analysis of head and neck carcinoma cells’ complex response to WEE1 inhibition has highlighted both the known and the previously uncharacterized deficiencies associated with p53 inactivation, and outlined specific directions for further inquiry and for therapeutic exploitation.

E. Mendez received commercial research grants and other commercial research support from AstraZeneca. No potential conflicts of interest were disclosed by the other authors.

Conception and design: A. Diab, M. Kao, K. Kehrli, J. Sidorova, E. Mendez

Development of methodology: A. Diab, M. Kao, J. Sidorova, E. Mendez

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): A. Diab, M. Kao, K. Kehrli, H.Y. Kim, J. Sidorova

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): A. Diab, M. Kao, J. Sidorova, E. Mendez

Writing, review, and/or revision of the manuscript: A. Diab, J. Sidorova, E. Mendez

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): H.Y. Kim

Study supervision: J. Sidorova, E. Mendez

This work was supported by the NIH/NCI grant R01 CA215647 to E. Mendez and J. Sidorova, Seattle Translational Tumor Research (STTR) programmatic investment grant to A. Diab, and University of Washington Royalty Research Fund pilot grant to J. Sidorova. This research was also supported by the Cellular Imaging and Therapeutic Manufacturing Shared Resources of the Fred Hutch/University of Washington Cancer Consortium (P30 CA015704).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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