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[Cancer Research 65, 7091-7095, August 15, 2005]
© 2005 American Association for Cancer Research

Priority Reports

Heterozygous ATR Mutations in Mismatch Repair–Deficient Cancer Cells Have Functional Significance

Kriste A. Lewis1, Sally Mullany1, Bijoy Thomas1, Jeremy Chien2, Ralitsa Loewen1, Viji Shridhar2 and William A. Cliby1

Departments of 1 Obstetrics and Gynecology and 2 Laboratory Medicine and Pathology, Mayo Clinic and Foundation, Rochester, Minnesota

Requests for reprints: William A. Cliby, Department of Obstetrics and Gynecology, Mayo Clinic and Foundation, 1317 Guggenheim, 200 1st Street, Southwest Rochester, MN 55905. Phone: 507-266-9323; Fax: 507-266-9300; E-mail: Cliby.william{at}mayo.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
ATR (ataxia telangiectasia and Rad3-related) function is necessary for the proper response to commonly used chemotherapeutic agents. Heterozygous truncating mutations in exon 10 of the ATR gene have been described in numerous cancers exhibiting microsatellite instability. We show that truncating mutations of ATR are capable of acting in a dominant-negative manner to abrogate ATR-dependent Chk1 phosphorylation and cell-cycle arrests after DNA damage. In addition, endometrial cell lines harboring ATR mutations are defective for ATR-dependent responses. These findings imply that ATR mutations play an important role in the development and clinical behavior of a subset of microsatellite instability–positive endometrial, colon, and stomach cancers.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Somatic mutations in exon 10 of ATR (ataxia telangiectasia and Rad3-related) occur in sporadic microsatellite instability (MSI)–positive stomach tumors (1). These mutations occur in a stretch of (A)10 repeats. Deletion or addition to (A)9 or (A)11, results in subsequent truncation at codon 778 or 776, respectively. Importantly, the wild-type (A)10 sequence has been identified in all samples tested. Subsequently, similar findings were reported for MSI+ endometrial hyperplasia and cancer (2). Given that ATR knock-out animals and cell lines are nonviable (3), and the long-term expression of kinase-inactive alleles of ATR is cytotoxic (4) the heterozygous nature of these mutations is likely necessary for survival. However, it also suggests that mutations in ATR in mismatch repair (MMR)–deficient cells may be important in the subsequent development and behavior of specific cancers. The association between inherited mutations in MMR genes (hereditary non–polyposis colon cancer) and the development of colon, stomach, and endometrial cancers makes further investigation of these findings critical. We present evidence that somatic truncating mutations in exon 10 of ATR are capable of acting in a dominant-negative manner and inhibit ATR-dependent responses to DNA damage.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell lines. HEC1A, AN3CA, Ishikawa, KLE, and RL95 endometrial cancer cell lines were provided by Dr. Paul Goodfellow (Washington University, St. Louis, MO): KLE is MMR-proficient and the remainder are MMR– and exhibit MSI (5). HCT116 and DLD1 (6, 7) are MMR– colon cancer cell lines; IGROV1 and SKOV3 are MMR– ovarian cancer cell lines (8). K562 is a human chronic myelogenous leukemia cell line.

Antibodies. Anti-human rabbit polyclonal ATR antibody was raised against amino acids 2,381 to 2,644 (Serotec, Oxford, England); ATR (N-19) goat polyclonal antibody was raised against the NH2 terminus of ATR; Chk1 mouse monoclonal antibody detects amino acids 1 to 476 of Chk1 protein (Santa Cruz Biotechnology, Santa Cruz, CA); Myc-Tag (9B11) mouse monoclonal antibody recognizes the myc-epitope; and anti-pChk1 polyclonal antibodies are phosphospecific antibodies against Ser317 and Ser345 (Cell Signaling Technology, Beverly, MA). Horseradish peroxidase–linked anti-rabbit, -goat, or -mouse antibodies were used (Cell Signaling Technology and Kirkegaard & Perry Laboratories, Gaithersburg, MA).

Mutational analysis. Nucleic acids were extracted using Qiagen (Valencia, CA) protocols. cDNA was prepared using Invitrogen (Carlsbad, CA) SuperScript first-strand synthesis system.

Primers complementary to flanking intronic sequences were designed to amplify the genomic fragment containing exon 10 (5'-CTAGGCTTTGTTTTACCAGT-3' forward, and 5'-TCAAGGCTTCAGTCTAATTC-3' reverse). To confirm if altered sequences were expressed, specific cDNA primers were used to amplify exon 10 and portions of exons 9 and 11 (5'-CCTTGAGTGGAGAACAGCAG-3' forward, and 5'-TCCATCTTCAGAGTCCAAGG-3' reverse). PCR was done using AmpliTaq Gold PCR Master Mix (Applied Bioscience, Foster City, CA) in a Bio-Rad (Hercules, CA) iCycler with annealing at 52°C (genomic DNA) or 56°C (cDNA). Products were sequenced using ABI PRISM Big Dye Terminator cycle sequencing ready reaction kit with AmpliTaq DNA Polymerase, FS version 1.1 and ABI PRISM 3730 DNA analyzer (96-capillary; Perkin-Elmer Applied Biosystems, Foster City, CA). Samples heterozygous for ATR alterations were subcloned in the topo-TA cloning vector, independently selected, and sequenced.

Construction of ATR mutant expression vector (ATR-stop). The complete coding sequence of wild-type ATR has the first ATG codon beginning at 106 bp, encoding a 2,464 amino acid protein, with a stop at codon 2,465. (A)9 deletion mutants result in a premature stop at codon 778 and (A)11 mutants at 776 (Fig. 1A). We constructed an ATR truncation expression vector (ATR-stop) containing 767 codons of ATR to perform functional studies simulating the deletion or insertion mutants. To allow specific immunodetection, a myc-tagged expression plasmid (pcDNA3.1/myc-HisA) was created with a 796 codon in-frame fusion protein with ATR-stop and the vector sequences for myc and His residues (Fig. 1A). Restriction sites were added to the primers for the enzymes BamHI (forward primer, 5'-GCGCGGATCCACTAGTGCCTCGCAGCCTCA-3') and Xho (reverse primer, 5'-GCGCCTCGAGAAGGAATGGCTTGCAGACAG-3'). Routine PCR was done under standard conditions for 30 cycles with annealing at 56°C. The resultant 2,344 bp fragment was then digested with BamHI and XhoI and cloned into similarly digested plasmid pcDNA3.1/myc-HisA using T4 DNA ligase. After ligation for 14 hours and transformation into DH5{alpha} competent cells, ampicillin-resistant colonies were screened by restriction digest, and the resultant ATR-stop vector was sequenced throughout the coding region.



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  		Figure 1. Exon 10 mutations in ATR. A, codons 766 to 779 and the corresponding wild-type sequence for exon 10 of ATR are shown. The (A)11 and (A)9 mutations are illustrated with the resultant premature stop codons. ATR-stop vector sequence is illustrated creating a fusion protein with myc-his sequences at the 3'-end of pcDNA3.1 myc-his. B, whole cell lysates were analyzed directly or immunoprecipitated using an NH2-terminal antibody and probed for ATR. Bands migrating at the expected size for ATR were seen in all endometrial cell lines. Faster migrating bands were observed in whole cell lysates for both Ishikawa and HEC1A cell lines. Samples immunoprecipitated with the NH2-terminal ATR antibody revealed a truncated form only in HEC1A cell line.

 
ATR expression and Chk1 activation assays. K562 cells were transiently transfected with empty vector (pcDNA3.1/myc-HisA) or ATR-stop plasmid DNA, incubated for 5 minutes and electroporated at 330 V for 5 ms x two pulses at low voltage in a T820 Electro Square Porator (BTX, Holliston, MA). Protein expression was confirmed by immunoblotting using antibodies to both ATR and myc-epitope. Cells for Western analysis were solubilized 30 minutes on ice in lysis buffer containing protease and phosphatase inhibitors. Samples were sonicated and centrifuged to pellet insolubles. Supernatant protein content was determined, lysate was combined with SDS-sample buffer, heated for 20 minutes at 68°C, separated on a 4% to 20% SDS-PAGE gel and electrophoretically transferred to nitrocellulose. Blots were probed (9) using reagents listed above. To examine native ATR protein expression, 400 µg of lysate were immunoprecipitated with protein G Sepharose-ATR (N-19) antibody complex. The immunoprecipitates were washed, eluted into SDS sample buffer, and heated for 20 minutes at 68°C before applying to a 6% SDS-PAGE gel for electrophoretic separation. The proteins were transferred to nitrocellulose and developed using Serotec rabbit anti-human ATR and anti-rabbit secondary antibody with enhanced chemiluminescence detection.

Flow cytometry. Control and treated cells were fixed in 50% ethanol, washed, digested with RNase A, stained with propidium iodide, and subjected to flow microfluorimetry on a FACScan flow cytometer (Becton Dickinson, Mountain View, CA) as previously described (9).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
ATR exon 10 mutations in endometrial and colon cancer cell lines. We found (A)10 mutations in MMR– endometrial cancer cell lines HEC1A and Ishikawa, but not in MMR– colon (HCT116 and DLD1) or ovarian cancer cell lines (IGROV1 and SKOV3). All mutations were heterozygous with wild-type (A)10 (Fig. 1A). Subcloning confirmed mutations were present in both genomic and cDNA. Either mutation leads to a frameshift with a stop codon (1), resulting in a truncated ATR lacking the carboxyl-terminal portion of the protein (Fig. 1A).

Basal levels of ATR vary within and among cell lines from different tissue types. Full-length ATR was observed in whole cell lysates and after immunoprecipitation in all endometrial cancer cell lines (Fig. 1B), although a decrease was noted in whole cell lysates from HEC1A and Ishikawa, consistent with haploinsufficiency anticipated by mutational inactivation of one allele. To confirm the presence of full-length ATR protein, immunoprecipitation was used to verify this observation (Fig. 1B). We noted faster migrating bands in HEC1A and Ishikawa cell lines using whole cell lysates probed with antibody raised against the NH2-terminal portion of ATR. These bands migrated at significantly lower molecular weight than would be anticipated based upon the predicted truncated protein and may therefore represent degradation products of the truncated protein as we would anticipate the posttranslational activity of the resultant protein to be markedly altered. To rule out the presence of additional mutations as a cause for these abnormally migrating forms, we have completely sequenced all 47 exons and intron-exon boundaries of ATR as part of an ongoing project to be reported.3 We observed no additional mutations in the coding sequences of the remainder of the endometrial cell lines (data not shown). The heterozygous mutations in cDNA and immunoblotting results suggest that both wild-type and mutated ATR are capable of being expressed at the protein level.

Transient expression of a truncated ATR protein abrogates damage-induced Chk1 phosphorylation and cell cycle arrest. Human K562 cells were transiently transfected with the ATR-stop plasmid. Expression of myc-tagged protein at the anticipated molecular weight 24 hours after transfection was confirmed (Fig. 2A). Both full-length and ATR-stop proteins also were detected using the N-19 anti-ATR antibody (Fig. 2E). Importantly, expression of ATR-stop did not seem to alter wild-type ATR levels. We consistently observed lower expression of ATR-stop relative to native wild-type ATR. Most of this difference could be ascribed to transfection efficiency (30%) in the K562 cells, however, we cannot rule out decreased protein stability for the truncated protein.



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  		Figure 2. Expression of a truncated ATR protein results in inhibition of Chk1 activation and cell cycle arrest seen after DNA damage. Expression of protein from the ATR-stop plasmid was confirmed using both anti-myc antibody (A) and an NH2-terminal anti-ATR antibody (E). Asynchronously growing K562 cells were transfected with control or ATR-stop plasmid and 24 hours later exposed to50 Jm–2 UV. After 1 hour, cell lysates were prepared and immunoblotted for expression of phosphospecific forms of Chk1 (B and C) or total Chk1 (D). Cells transfected with control plasmid showed strong phosphorylation of both Ser317 and Ser345 relative to untreated cells. F, K562 cells were transfected with ATR-stop plasmid or control parental plasmid or mock-transfected only. Twenty-four hours later, cells were harvested, or exposed to 125 nmol/L topotecan for 8 hours and analyzed. Marked S phase arrest is seen in control cells and cells transfected with parental control plasmid. This is abrogated in cells transfected with ATR-stop and is consistent with the Chk1 phosphorylation results.

 
We wanted to test whether ATR-stop expression abrogates UV-induced ATR-dependent Chk1 activation. Twenty-four hours after transient transfection with ATR-stop or empty parental vector, cells were exposed to 50 Jm–2 UV for 1 hour and lysates examined for Chk1 phosphorylation. Figure 2B and C show that cells transfected with empty vector were capable of strong Chk1 phosphorylation on Ser317 and Ser345: this effect was indistinguishable from untransfected controls (data not shown). However, in cells expressing ATR-stop, UV-induced Chk1 phosphorylation is diminished at both residues, and particularly at Ser345, a well-established target of ATR kinase activity. Given that wild-type ATR levels are not altered, this shows that expression of ATR-stop protein is capable of dominant-negative interference in downstream activation of Chk1, implying that such heterozygous mutations in cancers may be functionally relevant.

Similarly, we reasoned that the normally observed cell cycle arrest after S phase DNA damage might be blocked by ATR-stop expression. We have previously shown that prolonged exposure to the topoisomerase I poison topotecan results in S phase arrest that is ATR- and Chk1-dependent (10). Asynchronously growing K562 cells were transfected with either ATR-stop vector or empty vector (Fig. 2F). After 24 hours, cells were exposed to 125 nmol/L topotecan or diluent (control) for 24 hours and harvested for cell cycle analysis. As expected, K562 cells with empty vector showed a strong arrest in late S phase and G2. However, cells transiently transfected with ATR-stop failed to arrest. The observations that ATR-stop expression results in impaired damage-induced Chk1 phosphorylation and altered cell cycle arrest after S phase–specific DNA damage is consistent with the concept that cancer-associated ATR mutations are important in response to therapy.

Endometrial cancer cell lines with mutations in exon 10 of ATR are deficient in ATR-dependent DNA damage response. To initially test our in vitro observations, we examined the mutated endometrial cancer cell lines HEC1A and Ishikawa. Based on the above data, we expected to see evidence of altered Chk1 activation after DNA damage. Asynchronously growing cells were exposed to UV, ionizing radiation, or topotecan for the indicated time and concentrations (Fig. 3). We observed that K562 cells showed phosphorylation on Ser345 and Ser317 in response to all three damaging agents. However, the two endometrial cell lines were incapable of Ser345 phosphorylation after DNA damage, and we observed a decreased phosphorylation on Ser317 specifically after UV exposure. This was entirely consistent with the observed effects on Ser345 activation after UV damage in cells expressing ATR-stop (Fig. 2C).



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  		Figure 3. Endometrial cell lines with heterozygous ATR mutations in exon 10 have impaired ability to activate Chk1 after DNA damage. A, K562, HEC1A, and Ishikawa cells were treated with either diluent (DMSO control), 125 nmol/L topotecan for 8 hours, 10 Jm–2 UV for 2 hours, or 10 Gy IR for 2 hours. The cells were lysed, 50 µg of protein was applied to each lane, and following electrophoresis, the membranes were blotted using antibodies to specifically detect either total Chk1 or Chk1 phosphorylated on Ser317 or Ser345. ATR-dependent phosphorylation of Chk1-Ser345 is nearly absent after exposure to topotecan, UV, or IR relative to control cells. Ser317 phosphorylation seems to be partially maintained after topotecan and IR. B, cell cycle analysis showed loss of topotecan-induced S phase arrest in Ishikawa cells, but maintenance of arrest in HEC1A cells suggesting complex overlap in the S phase checkpoint response.

 
To examine cell cycle arrest after S phase damage, K562, HEC1A, and Ishikawa cell lines were exposed to 125 nmol/L topotecan or diluent for 24 hours. It should be noted that untreated asynchronously growing HEC1A cells consistently showed a markedly different cell cycle distribution compared with K562 or Ishikawa cells with a larger percentage of aneuploid cells (visible in Fig. 3B as a post-4n peak). Specifically, HEC1A cells show a higher percentage of S (39%) and G2 (41%) cell populations (Fig. 3B) versus K562 (26% and 36%, respectively) or Ishikawa (17% and 32%, respectively) cell lines: these steady-state differences make direct comparison of treated cell cycle results problematic using HEC1A cells but they are shown for completeness. The profile of untreated Ishikawa cells is similar to K562. Nontransfected K562 cells showed a predominant S phase arrest after 24 hours of exposure to topotecan (73% S phase versus 26% untreated) with few cells reaching the G2-M checkpoint. In contrast, Ishikawa cells show an abrogated S phase arrest and only a partial G2 arrest after damage that correlates with the observed inability to phosphorylate Ser345 after topotecan. A similar loss of S phase arrest was seen in the HEC1A cell lines (49% versus 39% untreated), although the abnormal untreated profile of these cells makes comparisons less clear. These results are consistent with our in vitro observations, and strongly suggest that ATR heterozygous mutations have in vivo consequences after DNA damage. These findings also imply that other pathways may partially compensate for ATR mutations.

Taken together, our results imply that ATR is a clinically relevant target of mutational inactivation in MMR-defective cancers. ATR mutations are likely to have important consequences in terms of clinical behavior and response to chemotherapy. Given the prevalence of MSI+ endometrial and colon cancers, our results suggest that clinical response to chemotherapeutic agents may be affected by the presence of such changes.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We show that MSI-associated frameshift mutations in ATR are capable of abrogating ATR-dependent DNA damage response. These findings are significant in the field of human cancer as they further add to the list of genes likely to be important mutational targets in MMR– cells and provide a potentially important link between mutations affecting ATR and sporadic human disease.

Our findings are consistent with the recent demonstration that haploinsufficiency of ATR can inhibit damage-induced phosphorylation of Ser345 (11). Additionally, Ser345 phosphorylation has been associated with chromatin association and binding of 14-3-3 protein (12). Future work will focus on the mechanisms by which truncating ATR mutations block specific aspects of DNA damage response. Such studies are likely to shed meaningful light on the specifics of ATR activation, and the alternative compensatory pathways in such cancers. This is also consistent with the paradigm that MSI-associated mutations in important members of the DNA damage response system are functionally relevant. Such alterations have been described for Mre11 in MSI+ stomach cancers as well as the colon cancer cell line HCT116 (13, 14). Notably, functional Mre11 mutations occurred in introns, as opposed to exonic mutations described for ATR to date. The observation that defects in MMR results in selective alterations of certain components of the DNA damage response implies a selection bias and suggests synthetic lethality for some alterations.

The timing of ATR mutations in the development or progression of MSI+ cancers is unknown. Several lines of evidence make it likely that ATR alterations are important: (a) the well-described role of ATR in checkpoint control (15, 16), (b) the proposed role for ATR in maintaining genomic stability (17), (c) the present data demonstrating impaired DNA damage response in cancer harboring exon 10 mutations, and (d) the recent report that ATR acts as a dosage-dependent tumor suppressor in MMR-deficient background (11). Heterozygous mutations in ATR have been described in primary endometrial and stomach cancers (1, 2, 18). We did not observe exon 10 mutations in the colon or ovarian cancer cell lines that we screened.

Recent studies have described a possible interaction between ATR and MMR proteins (19, 20). In light of the current data, it will be important to confirm the mutational status of ATR in such experiments to avoid erroneously ascribing these observations to an interaction between ATR and MMR proteins as opposed to an existing inactivating ATR mutation.

We have previously shown that ATR kinase–defective cells are hypersensitive to cisplatin and topoisomerase inhibitors. As both agents are commonly used to treat gynecologic malignancies, future studies will be needed to assess the impact of such alterations on the response to these agents. Importantly, ATR mutations in MSI+ cancers afford a potential mechanism of achieving a therapeutic advantage relative to normal tissues. Alternatively, such cancers may be particularly susceptible to further inhibition of ATR or Chk1.


   Acknowledgments
 
Grant support: National Cancer Institute grant CA87579 (to W.A. Cliby).

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.


   Footnotes
 
3 Manuscript in preparation. Back

Received 3/28/05. Revised 5/31/05. Accepted 6/ 9/05.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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  10. Cliby WA, Lewis KA, Lilly KK, Kaufmann SH. S phase and G2 arrests induced by topoisomerase I poisons are dependent on ATR kinase function. J Biol Chem 2002;277:1599–606.[Abstract/Free Full Text]
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