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Cancer Research 68, 7947, October 1, 2008. doi: 10.1158/0008-5472.CAN-08-0971
© 2008 American Association for Cancer Research
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Experimental Therapeutics, Molecular Targets, and Chemical Biology

ATM and the Mre11-Rad50-Nbs1 Complex Respond to Nucleoside Analogue–Induced Stalled Replication Forks and Contribute to Drug Resistance

Brett Ewald, Deepa Sampath and William Plunkett

Department of Experimental Therapeutics, The University of Texas M. D. Anderson Cancer Center and The University of Texas Health Science Center at Houston Graduate School of Biomedical Sciences, Houston, Texas

Requests for reprints: William Plunkett, Department of Experimental Therapeutics, Unit 71, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Boulevard, Houston, TX 77030. Phone: 713-792-3335; Fax: 713-794-4316; E-mail: wplunket{at}mdanderson.org.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 References
 
The Mre11-Rad50-Nbs1 complex and autophosphorylated Ser1981-ATM are involved in recognizing and repairing DNA damage, such as double-strand breaks (DSB). However, the role of these factors in response to stalled replication forks is not clear. Nucleoside analogues are agents that are incorporated into DNA during replication, which cause stalling of replication forks. The molecular mechanisms that sense these events may signal for DNA repair and contribute to survival but are poorly understood. Cellular responses to both DSBs and stalled replication forks are marked by H2AX phosphorylation on Ser139 ({gamma}-H2AX), which forms nuclear foci at sites of DNA damage. Here, concentrations of the nucleoside analogues 1-β-D-arabinofuranosylcytosine (cytarabine; ara-C), gemcitabine, and troxacitabine, which inhibited DNA synthesis by 90% within 2 hours, were determined for each agent. Using {gamma}-H2AX as a marker for changes in chromatin structure, we show that Mre11, Rad50, Nbs1, and phosphorylated ATM respond to nucleoside analogue–induced stalled replication forks by forming nuclear foci that colocalize with {gamma}-H2AX within 2 hours. Because neither DSBs nor single-strand breaks were detectable after nucleoside analogue exposure, we conclude that this molecular response is not due to the presence of DNA breaks. Deficiencies in ATM, Mre11, or Rad50 led to a 2- to 5-fold increase in clonogenic sensitization to gemcitabine, whereas Nbs1 and H2AX deficiency did not affect reproductive growth. Taken together, these results suggest that ATM, Mre11, and Rad50 are required for survival after replication fork stalling, whereas Nbs1 and H2AX are inconsequential. [Cancer Res 2008;68(19):7947–55]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 References
 
Endogenous and exogenous events that hinder replication fork progression threaten both cellular survival and genome integrity. Therefore, molecular mechanisms that monitor and regulate fork progression are necessary to safeguard DNA replication under these circumstances (1). However, this capability likely contributes to resistance to pharmacologic agents that target DNA replication, such as those that cause steric hindrance (i.e., aphidicolin), decrease the deoxynucleotide pool (i.e., hydroxyurea), or block DNA synthesis of the nascent strand (i.e., nucleoside analogues). Upon cellular entry and phosphorylation to their active metabolites, many nucleoside analogues are incorporated into replicating DNA and cause replication forks to stall, as they are poor substrates for chain extension (2). Cells respond to these events by activating the ATR-Chk1-Cdk2 S-phase checkpoint pathway (3, 4), which blocks firing of new replication origins and enforces cell cycle arrest. This likely enables DNA repair mechanisms to react to sites of DNA damage and contribute to survival.

The molecules that recognize nucleoside analogue–induced stalled replication forks and signal for repair processes are not well-characterized. Our prior studies showed that cells respond rapidly to nucleoside analogue exposure by causing phosphorylation of the DNA damage responsive histone, H2AX, and the phosphoinositol kinase–like kinase, ATM (4). The function of H2AX is not fully understood, but it is likely involved in anchoring damage-response proteins to sites of DNA damage, whereas ATM is a regulator of DNA repair and cell cycle checkpoints (5). Because phosphorylation of H2AX and ATM are most closely associated with recognition of DNA double-strand breaks (DSB; ref. 6), it was not expected that such pronounced responses would occur upon nucleoside analogue exposure. It is now well-established that H2AX is rapidly phosphorylated in response to additional forms of DNA damage, such as single-stand breaks (SSB; ref. 7), stalled replication forks (4, 8), and hyperthermia-induced stresses (9). The molecules that are associated with the recognition and repair of ionizing radiation–induced DNA damage have been well-characterized and may offer insight into cellular responses to blocked DNA replication.

The Mre11-Rad50-Nbs1 (MRN) complex is involved in the cellular responses to DSBs and may also function during normal DNA replication (10, 11). It is among the first responders to bind to DNA ends (12, 13) and has been shown to tether DNA fragments through the self-association of Rad50 coiled-coil domains (14). Mre11 possesses 3'->5' exonuclease and single-strand endonuclease activities (15, 16) and, therefore, potentially has the capability to remove nucleoside analogues from DNA. The DNA damage response is a highly regulated concert of many molecules in which the MRN complex functions both upstream and downstream of ATM. Here, we examined molecules that are known regulators of the DSB response to determine if they recognize nucleoside analogue–induced stalled replication forks and contribute to survival.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosure of Potential...
 References
 
Cell culture. The following cell lines were used: OCI-AML3 adult myelogenous leukemia cell line (gift from M. Andreeff, The University of Texas M. D. Anderson Cancer Center, Houston, TX) was maintained in RPMI 1640 supplemented with 10% heat-inactivated fetal bovine serum (FBS; Invitrogen). Ataxia-telangiectasia (AT) fibroblasts, AT22IJE-T+vector, and those that have been repleted with full-length ATM (AT22IJE-T+ATM, gifts from Y. Shiloh, Sackler School of Medicine, Tel Aviv, Israel; ref. 17) were maintained in DMEM plus 20% FBS. Nijmegen breakage syndrome (NBS) fibroblasts, GM7166VA7, and those repleted with full-length Nbs1 (GM7166VA7+Nbs1, gifts from K. Komatsu, Kyoto University, Kyoto, Japan; ref. 18), and H2AX–/– and H2AX+/+ mouse embryonic fibroblasts (gifts from A. Nussenzweig, National Cancer Institute, Bethesda, MD; ref. 19) were maintained in DMEM plus 15% FBS.

Chemicals and antibodies. The nucleoside analogues, gemcitabine and troxacitabine, were kindly provided by Dr. L. W. Hertel (Lilly Research Laboratories, Indianapolis, IN) and Dr. Henriette Gourdeau (Biochem Pharma, Montreal, Canada), respectively. The 1-β-D-arabinofuranosylcytosine (ara-C) used in these investigations was purchased from Sigma-Aldrich. Mouse monoclonal antibodies used were purchased from Upstate Biotechnology (pSer139-H2AX, pSer1981-ATM), Abcam (ATM), Novus Biologicals (Mre11), GeneTex (Rad50), Lab Vision (Ku70, Ku80), and Sigma-Aldrich (β-actin). Rabbit polyclonal antibodies were purchased from Bethyl Laboratories (pSer139-H2AX) and Novus Biologicals (Nbs1). Alexa Fluor 488, 594, and 680 fluorescent secondary antibodies were purchased from Molecular Probes.

Immunoblotting analysis. Cell lysate preparations and detection of specific proteins were done according to previously described procedures (4). The Odyssey IR imaging system (Li-Cor Biosciences) was used according to the manufacturer's instructions for protein visualization.

DNA synthesis assay. Measurement of DNA synthesis was done as previously described (4). Briefly, [3H]thymidine (5 µCi/mL, 60.9 Ci/mmol; Moravek Biochemicals) was added 30 min before harvesting. Labeled cells were collected on glass fiber filters (Schleicher & Scheull), and radioactivity retained on the filter was determined by a liquid scintillation counter (Packard).

Confocal microscopy. OCI-AML3 cultures were centrifuged onto slides, fixed, and immunostained according to previously described procedures (4). Images were obtained using an Olympus FluoView IX71 confocal microscope system (Olympus) and processed using FluoView FV500 ver. 5 software. The objective lens was a x40/1.30 numerical aperature oil lens. A profile of overlapping red and green fluorescence was measured by the image processing and analysis software, ImageJ (NIH). Pearson's correlation coefficient (r) was calculated for each image using the Olympus FluoView FV500 ver. 5 software.

Cell cycle analysis. Cells were washed with ice-cold PBS, fixed, and stained with propidium iodide as previously described (4). At least 10,000 cells were evaluated for fluorescence using a Becton Dickinson FACSCalibur flow cytometer.

Comet assay. OCI-AML3 cells were mixed with agarose, placed onto a microscope slide, and assessed for DNA damage, as previously described (20). Briefly, the slides were left in lysis solution for 1 h at 4°C in the dark, washed twice with 1x Tris-borate EDTA buffer (neutral conditions) or incubated in alkaline solution (pH >13) for 1 h, and subjected to electrophoresis (300 mA) for 15 min at 4°C. Comet images analyzed by a computer-based image analysis system (Kinetic Imaging Komet system, Version 5.5) for tail moment, DNA content, and percentage of DNA in the tail.

Pulsed-field gel electrophoresis. Exponentially growing cultures were prelabeled with [14C]thymidine for 48 h (0.02 µCi/mL every 12 h x 4), exposed to drugs or ionizing radiation (3.17 Gy/min of 137Cs), and harvested immediately. Cells were washed, embedded in agarose plugs, lysed, digested with proteinase K, and the DNA was separated by pulsed-field gel electrophoresis (PFGE) using a CHEF-DR III system (Bio-Rad Laboratories), as previously described (21). The gel was stained with ethidium bromide and imaged using the ImageQuant software program (Molecular Dynamics). Radioactivity was quantitated using the InstantImager system (PerkinElmer) after the gel had been dried at 60°C under vacuum.

siRNA transfections. AT22IJE-T+ATM cells were plated in 6-well plates and incubated overnight. Medium was replaced with fresh medium containing 100 nmol/L siControl nontargeting siRNA (Dharmacon), Mre11 siRNA (Dharmacon), Rad50 siRNA (Dharmacon), or Nbs1 siRNA (Dharmacon) and DharmaFECT 1 transfection reagent (Dharmacon). Seventy-two hours after transfection, cells were harvested and plated for clonogenic survival assays. The remaining cells were lysed and subsequently examined for protein knockdown by immunoblotting.

Clonogenic assays. Exponentially growing cultures were placed in a 6-well plate for 8 to 24 h before being exposed to the indicated drugs for 1 cell cycle period. Cells were then washed twice with PBS (37°C), fresh medium was added, and allowed to grow undisturbed for 7 to 10 d in normal growing conditions. Cells were fixed (50% methanol, 5% acetic acid, 45% H2O) for 1 h, stained with Accustain Giemsa (1:20 dilution; Sigma-Aldrich) for 1 h, and washed twice with water. Colonies of ≥50 cells were counted using a dissecting microscope.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosure of Potential...
 References
 
Inhibition of DNA synthesis after exposure to nucleoside analogues. The deoxycytidine nucleoside analogues, cytarabine (ara-C), gemcitabine, and troxacitabine (Fig. 1A ), were used to cause stalled replication forks in the acute myelogenous leukemia cell line, OCI-AML3. A range of nucleoside analogue concentrations was first investigated to determine what concentrations of each drug caused a similar inhibition of DNA replication (data not shown). Exposure of 0.5 µmol/L ara-C, 0.1 µmol/L gemcitabine, or 2 µmol/L troxacitabine caused inhibition of DNA synthesis by ~70% within 1 h and 90% within 2 h, as measured by a decrease in [3H]thymidine incorporation (Fig. 1B). The decrease in DNA replication caused by drug incorporation into DNA was associated with an accumulation of cells in S phase (>60%) within 24 h, as determined by propidium iodide staining of DNA content (Fig. 1C). These investigations show that the deoxycytidine nucleoside analogues ara-C, gemcitabine, and troxacitabine are incorporated into DNA, causing an early inhibition of DNA synthesis and eventual arrest in S phase.


Figure 1
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  		Figure 1. The effect of nucleoside analogues on DNA synthesis. A, chemical structures of (i) 2'-deoxycytidine, (ii) ara-C, (iii) gemcitabine, and (iv) troxacitabine. B, effect of 0.5 µmol/L ara-C (bullet), 0.1 µmol/L gemcitabine ({circ}), or 2 µmol/L troxacitabine ({blacktriangleup}) on DNA synthesis in OCI-AML3 cultures, as measured by [3H]thymidine incorporation after a 30-min pulse. Points, mean; bars, SD. C, accumulation of OCI-AML3 cells in early S phase after exposure to 1 µmol/L ara-C, 10 nmol/L gemcitabine, or 250 nmol/L troxacitabine for 24 h, as measured by DNA content (propidium iodide). Cell populations representing G1, S, and G2-M phases are appropriately marked.

 
Colocalization of phosphorylated ATM and the MRN complex at sites of nucleoside analogue–induced DNA damage. The histone variant, H2AX, is rapidly phosphorylated and forms distinct nuclear foci in response to many forms of DNA damage, such as DSBs (22) and stalled replication forks (4, 8). We investigated if other molecules that are important for the sensing and repairing of DNA DSBs also accumulate at sites of nucleoside analogue–induced stalled replication forks. Short drug incubations were chosen to minimize the effect of DSBs possibly caused by the collapse of replication forks. OCI-AML3 cultures exposed to ara-C, gemcitabine, and troxacitabine, which caused an inhibition of DNA synthesis by 90% within 2 h (Fig. 1B), showed an increase in phosphorylation of Ser1981 on ATM (Fig. 2A ). Distinct pSer1981-ATM foci, which colocalized with {gamma}-H2AX, were evident within 2 hours of exposure to all three nucleoside analogues, as analyzed by confocal microscopy. Red-green profiles illustrate overlapping nuclear foci, as depicted by overlapping red and green fluorescence peaks, for a region in which a line was randomly drawn through the center of a nucleus (Fig. 2A, bottom). Pearson's correlation coefficient (r) was calculated for each image to determine the relationship between phosphorylated ATM and {gamma}-H2AX, where one represents 100% correlation and zero for no correlation (Table 1 ; ref. 23). The correlation of pSer1981-ATM and {gamma}-H2AX significantly (P < 0.001) increased from 30% to 61% to 71% after nucleoside analogue exposure, which was comparable with that found in cultures treated with 10 Gy of ionizing radiation (74%; Table 1).


Figure 2
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  		Figure 2. Colocalization of phosphorylated Ser1981-ATM, Rad50, Nbs1, and Mre11 at sites of stalled replication forks. Exponentially growing OCI-AML3 cultures were incubated with 0.5 µmol/L ara-C, 0.1 µmol/L gemcitabine (gem), or 2 µmol/L troxacitabine (trox) for 2 h or exposed to 10 Gy ionizing radiation (IR), harvested, and subjected to fluorescent staining. Nuclei were stained with 4',6-diamidino-2-phenylindole (DAPI) and are shown in the top in blue. A single line was randomly drawn through the center of nuclei to generate a red and green fluorescence profile (bottom). Overlapping peaks illustrate overlapping red/green nuclear foci. Representative images of two independent experiments taken by confocal microscopy of (A) pSer1981-ATM (green)/{gamma}-H2AX (red), (B) Rad50 (green)/{gamma}-H2AX (red), (C) Nbs1 (red)/{gamma}-H2AX (green), and (D) Mre11 (green)/{gamma}-H2AX (red) are shown.

 

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  		Table 1. Pearson's correlation coefficients of DNA damage response proteins with {gamma}-H2AX

 
Further investigations showed that Mre11, Rad50, and Nbs1 form nuclear foci in OCI-AML3 nuclei that also colocalize at sites of stalled replication forks within 2 hours of exposure to nucleoside analogues (Fig. 2B–D). An increased association of Rad50 and Nbs1 with {gamma}-H2AX was observed after exposure to these agents or ionizing radiation, as determined by increased colocalization of foci and Pearson's coefficient (Fig. 2B–C; Table 1). DNA damage–associated foci containing Mre11 and {gamma}-H2AX also increased after exposure nucleoside analogues or ionizing radiation, suggesting an Mre11 response (Fig. 2D). High levels of Mre11-associated fluorescence was evident throughout the nuclei of untreated cells (Fig. 2D, green) and likely explains a lack of an increase in Pearson's correlation coefficient between Mre11 and {gamma}-H2AX under all conditions (Table 1). DNA damage–induced nuclear foci were found in only a fraction of cells exposed to nucleoside analogues (Fig. 2; data not shown), which is likely because these agents cause stalled replication forks in cells actively undergoing DNA replication (~35% of total cell population). Conversely, an increase in DNA damage foci was evident in a majority of cells after ionizing radiation treatment, which is not cell cycle specific (Fig. 2A–D, right). A similar response was previously reported for {gamma}-H2AX (4). Although ara-C, gemcitabine, and troxacitabine have slightly different overall mechanisms of action (2), differences in colocalization was not dependent on the agent used. These experiments show that molecules that are closely associated with the DSB response also colocalize at sites of nucleoside analogue–induced stalled replication forks.

Undetectable levels of DNA breaks caused by nucleoside analogues. We investigated if the molecular response to nucleoside analogue–induced DNA damage was associated with the presence of DSBs because the functions of ATM, H2AX, and the MRN complex are most closely associated with the sensing, signaling, and repair of these lesions. OCI-AML3 cultures were exposed to nucleoside analogues or ionizing radiation and examined for the presence of DSBs by pulsed-field gel electrophoresis (PFGE). Cultures exposed to nucleoside analogues for 2 hours showed similar levels of DSB induction compared with untreated controls, as determined by both fluorescence (Fig. 3A, top ) and [14C]-labeling of DNA (Fig. 3A, bottom). Conversely, 10 to 40 Gy of ionizing radiation caused a 1.5- to 4-fold increase in DSB detection. Because the number of DSBs caused by 10 Gy of ionizing radiation approached the lower limit of detection using PFGE, cells were further examined by the comet assay under both neutral and alkaline conditions, as a potentially more sensitive method to measure DSBs and SSBs, respectively. Similar to PFGE experiments, OCI-AML3 cultures exposed to nucleoside analogues for 2 hours did not show increased levels of either DSBs (Fig. 3B) or SSBs (Fig. 3C), compared with untreated controls. Conversely, 10 Gy of ionizing radiation caused an increase in mean tail moment of 2- to 3-fold (neutral conditions) and 13-fold (alkaline conditions), which likely represents the measurement of DNA damage caused by ~400 DSBs per cell (24). The limit of sensitivity for the comet assay has been estimated at ~50 strand breaks per diploid mammalian cell (25). As there are opportunities for nucleoside analogues to incorporate into an estimated 100,000 replication forks throughout S phase, in which 10% are engaged in DNA synthesis at any one time (26, 27), it does not seem likely that the molecular response seen under these conditions (Fig. 2) is due to the presence of DNA breaks.


Figure 3
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  		Figure 3. Undetectable levels of DNA breaks after nucleoside analogue exposure. OCI-AML3 cultures were incubated with 0.1 µmol/L gemcitabine, 0.5 µmol/L ara-C, or 2 µmol/L troxacitabine for 2 h or exposed to 10 to 40 Gy ionizing radiation, harvested, and examined for DNA break induction. A, representative gel image of DNA stained with ethidium bromide in a typical experiment after PFGE (top). The lower band represents the fraction of DNA with DSBs. The DNA in the lower band divided by the total DNA in each lane determined by [14C] labeling, compared with untreated controls is graphed. Columns, mean for two independent experiments performed in duplicate; bars, SE. B and C, calculated mean tail moment after exposure to nucleoside analogues or ionizing radiation, as determined by comet assay under neutral (B) or alkaline (C) conditions. Columns, mean for two independent experiments in which 100 cells per experiment were analyzed; bars, SD. *, P < 0.001.

 
The contribution of DNA damage sensor and repair molecules to survival. ATM, H2AX, Mre11, Rad50, and Nbs1 play important roles in sensing and repairing DSBs (10). Deficiencies in these DNA damage response molecules cause increased sensitivity to ionizing radiation (19, 28–31), implying that their functions relate to survival with regard to DSBs. To determine if these proteins contribute to survival after the formation of stalled replication forks, we analyzed cell lines that are deficient or have decreased levels of ATM, Mre11, Rad50, Nbs1, or H2AX for their sensitivity to a nucleoside analogue. When immortalized AT fibroblasts deficient for ATM (Fig. 4A ) were exposed to gemcitabine, a significant difference in reproductive viability (P < 0.001) was observed, compared with the repleted line. Exposure of 10 nmol/L gemcitabine for 24 hours before drug removal killed 70% of AT cells, whereas <10% of ATM-repleted cells lost repopulation capacity (Fig. 4A, middle). A 4.8-fold difference in gemcitabine sensitivity was observed between AT and AT+ATM cultures after exposure to a range of gemcitabine concentrations, as determined by IC50 values (Fig. 4A, bottom). These results are consistent with a previous report that showed a 2-fold increase in sensitivity to gemcitabine after ATM knockdown using siRNA in A549 and HeLa cells (32).


Figure 4
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  		Figure 4. Effect of ATM, Mre11, and Rad50 on colony growth after exposure to gemcitabine. A, exponentially growing AT and AT+ATM fibroblasts were analyzed for ATM protein level by immunoblotting of whole cell lysates (top), exposed to 10 nmol/L gemcitabine for 24 h (middle) or a range of gemcitabine concentrations (bottom) before analysis of clonogenic survival after 10 d. B to D, exponentially growing AT+ATM cultures were untreated, transfected with control siRNA, Mre11 siRNA, or Rad50 siRNA. Seventy-two hours after transfection, cells were harvested and split into two samples. One sample was lysed and saved for immunoblotting (B). Cells from the other sample were replated and allowed to grow undisturbed for 8 h before being exposed to 10 nmol/L gemcitabine (B, bottom) or 1 to 100 nmol/L gemcitabine (C and D) for 24 h. Then, gemcitabine was washed out of the medium and colonies of untreated ({diamondsuit}), control siRNA (bullet), and Mre11/Rad50 siRNA ({circ}) were counted after 10 d of undisturbed growth. The mean of two independent experiments performed in triplicate (n = 6) ± SE is graphed. *, P < 0.001. Gem, 10 nmol/L gemcitabine.

 
In experiments in which Mre11 (85–90% knockdown; Fig. 4B, top; Supplementary Fig. S1) or Rad50 (75–78% knockdown; Fig. 4B, middle) was depleted in the AT+ATM cell line using targeted siRNA, a 35% to 50% decrease in clonogenic survival was observed when cells were exposed to 10 nmol/L gemcitabine, compared with siRNA controls (P < 0.001; Fig. 4B, bottom). When exposed to a range of gemcitabine concentrations, an approximate 2-fold increase in gemcitabine sensitivity was calculated in Mre11- and Rad50-depleted cells, as determined by calculated IC50 values (Fig. 4C–D). Consistent with other reports (33, 34), a decrease in Mre11 protein caused by siRNA also led to a reduction in Rad50 (85%) and Nbs1 (40%), suggesting that Mre11 contributes to Rad50 and Nbs1 protein stability (Fig. 4B; Supplementary Fig. S1). Decreases in Mre11 did not affect protein levels of other DNA damage sensors, Ku70 or Ku80 (Fig. 4B, top). Neither the Mre11 nor the Nbs1 protein was affected after knockdown of Rad50 (data not shown).

Immortalized NBS fibroblasts, which lack Nbs1 but were proficient for Mre11 and Rad50, did not show a difference in gemcitabine sensitivity, compared with Nbs1-repleted cultures (Fig. 5A ). The lack of increased sensitivity was confirmed by using Nbs1-targeted siRNA in AT+ATM cultures (Fig. 5B). Although Nbs1 has been reported to be required for the S-phase checkpoint in response to UV irradiation, ionizing radiation, and hydroxyurea (35, 36), Nbs1-deficient cells exposed to nucleoside analogues arrested in S phase at similar concentrations as the paired Nbs1-proficient cell line (Supplementary Fig. S2). Similarly, immortalized H2AX–/– mouse embryonic fibroblasts were not sensitized to gemcitabine (Fig. 5C). This result supports previous data that the DNA damage response after nucleoside analogue exposure was not due to the presence of DSBs (Fig. 3), as an increase in sensitivity is expected under DSB conditions (19). Figure 5D summarizes the IC50 values, 95% confidence intervals, and fold differences in gemcitabine sensitivity for all cell lines. This investigation shows that ATM, Mre11, and Rad50, but not Nbs1 and H2AX, contribute to cell survival and recovery from gemcitabine-induced stalled replication forks.


Figure 5
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  		Figure 5. Effect of DNA damage response molecules on colony growth after exposure to gemcitabine. Exponentially growing (A) NBS ({circ}) and NBS+Nbs1 (bullet) human fibroblasts, (B) AT+ATM human fibroblasts that were untreated ({diamondsuit}), transfected with control siRNA (bullet), or exposed to Nbs1 siRNA ({circ})before drug exposure, and (C) H2AX–/– ({circ}) and H2AX+/+ (bullet) mouse embryonic fibroblasts were exposed to a range of gemcitabine concentrations for one cell cycle (15–24 h) before fresh medium replacement. Cells were allowed to grow undisturbed for 5 to 10 d in normal growing conditions before colonies of ≥50 cells were counted. Mean of two independent experiments performed in triplicate are shown. Points, mean; bars, SE. IR, 10 Gy ionizing radiation. D, IC50 values, 95% confidence intervals (CI), and fold differences between compared conditions is summarized.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosure of Potential...
 References
 
ATM and the MRN complex are critical to sensing DSBs and initiating repair of damaged DNA. They may also function during DNA replication (11). Here, we report that these molecules respond to stalled replication forks, and that ATM, Mre11, and Rad50 are required for survival after such agitation. A previous study, showed a similar increase in sensitivity after depletion of ATR, which is a known regulator of the DNA damage response to stalled replication forks (32). However, the relationship between ATR and the DNA damage response molecules investigated here is not well-understood. Nbs1 and H2AX are not likely critically involved in the response to stalled replication forks because their absence did not affect reproductive viability (Fig. 5). It has been reported that Nbs1 is required for the S and G2-M cell cycle checkpoints in response to UV irradiation, hydroxyurea, or ionizing radiation (35, 36). However, we did not find that Nbs1 was required for a nucleoside analogue–induced S-phase arrest (Supplementary Fig. S2). Therefore, the molecular sensors investigated here likely have damage-specific functions and contribute to cell survival after inhibition of DNA synthesis by a different mechanism than signaling for cell cycle arrest. Differences may be due to the mechanisms that cause stalled replication forks. Nucleoside analogues physically block DNA synthesis of extending DNA strands, whereas UV and hydroxyurea cause fork stalling by other means. Because Nbs1 was not required for survival under these conditions, it is not clear if Mre11, Rad50, and Nbs1 function as a complex, as they do after DSBs, or individually in response to stalled replication forks.

A variety of human disorders are associated with depleted ATM, Mre11, or Nbs1, including AT, AT-like disorder, and NBS (5, 34, 37, 38). Patients with these disorders display similar characteristics of cancer susceptibility, extreme sensitivity to ionizing radiation, and neurologic defects. These phenotypes indicate that ATM, Mre11, and Nbs1 may play a role in normal DNA replication (11). In support of this hypothesis, it has been shown that Mre11 is required to prevent DSBs during normal chromosomal replication in Xenopus eggs extract (39). Furthermore, Mre11 associates with chromatin in an S phase–specific manner and localizes to sites of ongoing DNA replication in human fibroblasts, as visualized by proliferating cell nuclear antigen and incorporated bromodeoxyuridine (40). The results reported here extend the functions of ATM, Mre11, and Rad50 to promoting cell survival in response to blocked DNA extension of nascent strands.

Although evidence supports that nucleoside analogues incorporate into DNA and exert a cytotoxic effect by blocking DNA synthesis (Fig. 1; ref. 2), we sought to confirm that the molecular response observed here was not due to the presence of DNA breaks. Although the assays were sensitive enough to detect DSBs caused by ionizing radiation, these lesions were not measurable after nucleoside analogue exposure (Fig. 3). It is estimated that 1 Gy of ionizing radiation causes 40 DSBs per cell, resulting in a similar number of {gamma}-H2AX nuclear foci (24). The same 1:1 ratio is not likely true in response to agents that cause stalled replication forks. An estimated 50,000 replication origins containing two forks each are brought together to form many replication factories at the start of replication in human cells (41, 42). Therefore, it is likely that the number of DNA damage nuclear foci observed here represent replication factories containing many stalled replication forks. These factories are stably anchored in the nucleus but changes in the patterns occur during coordinated assembly and disassembly throughout S phase (43). The pattern that forms after nucleoside analogue exposure resembles similar patterns reported previously during early S phase DNA replication (Fig. 2; refs. 44, 45), supporting our earlier finding that H2AX phosphorylation first occurs after nucleoside analogue treatment in cells initiating DNA synthesis (4).

The DNA repair mechanisms involved in removing nucleoside analogues from DNA are not well-understood. Proof-reading 3'->5' exonuclease activities associated with replicative DNA polymerases (46) and base excision repair processes (47) are capable of removing fraudulent nucleotides from DNA. However, a slow rate of drug removal (48) and sustained cell cycle arrest after exposure to nucleoside analogues (49) suggests that these mechanisms do not significantly promote survival. It is unknown what role ATM, Mre11, and Rad50 play at replication forks when DNA synthesis is inhibited. These molecules may have replication fork surveillance or DNA repair mechanisms that respond to chemotherapeutic agent–induced replication blocking and, thus, contribute to drug resistance. Because Mre11 possesses 3'->5' exonuclease and single strand endonuclease activities (15, 16), it may have the capacity to remove nucleoside analogues from DNA. Excision kinetics are enhanced when Mre11 is coupled with Rad50 (15), which may offer an explanation for the increase in gemcitabine sensitivity observed in Rad50-depleted cultures (Fig. 4). Simplified models using purified enzymes/enzyme complexes, oligonucleotides, and primer extension assays may be useful to answer these questions and may uncover a novel mechanism for the removal of nucleoside analogues from DNA, which is currently poorly understood (50).

Here, we present mechanistic evidence for the importance of a DNA repair pathway not previously known to be required for survival after the induction of stalled replication forks. These results provide a rationale for further studies to determine the mechanisms by which these molecules contribute to survival.


   Disclosure of Potential Conflicts of Interest
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
References
 
No potential conflicts of interest were disclosed.


   Acknowledgments
 
Grant support: CA32839, CA55164, and Cancer Center support grant CA16672 from the National Cancer Institute, Department of Health and Human Services.

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.

We thank Dr. Raymond Meyn and Colin Brooks for their assistance with PFGE investigations and Dr. V. Ashutosh Rao for his assistance with confocal microscopy data analysis.


   Footnotes
 
Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

Received 3/13/08. Revised 6/14/08. Accepted 7/23/08.


   References
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 References
 

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