|Home | About | Journals | Submit | Contact Us | Français|
In this study, we examine the telomeric functions of the mammalian Mre11 complex by using hypomorphic Mre11 and Nbs1 mutants (Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB, respectively). No telomere shortening was observed in Mre11ATLD1/ATLD1 cells after extensive passage through culture, and the rate of telomere shortening in telomerase-deficient (TertΔ/Δ) Mre11ATLD1/ATLD1 cells was the same as that in TertΔ/Δ alone. Although telomeres from late-passage Mre11ATLD1/ATLD1 TertΔ/Δ cells were as short as those from TertΔ/Δ, the incidence of telomere fusions was reduced. This effect on fusions was also evident upon acute telomere dysfunction in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells rendered Trf2 deficient by cre-mediated TRF2 inactivation than in wild-type cells. The residual fusions formed in Mre11 complex mutant cells exhibited a strong tendency toward chromatid fusions, with an almost complete bias for fusion of telomeres replicated by the leading strand. Finally, the response to acute telomere dysfunction was strongly impaired by Mre11 complex hypomorphism, as the formation of telomere dysfunction-induced DNA damage foci was reduced in both cre-infected Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/F cells. These data indicate that the Mre11 complex influences the cellular response to telomere dysfunction, reminiscent of its influence on the response to interstitial DNA breaks, and suggest that it may promote telomeric DNA end processing during DNA replication.
The Mre11 complex (in mammals, Mre11, Rad50, and Nbs1) plays a central role in the cellular response to DNA double-strand breaks (DSBs). The Mre11 complex acts as a DSB sensor, promoting the activation of ATM-dependent DNA damage signaling pathways, DNA repair, and apoptosis. In addition, the complex plays a direct role in recombinational DNA repair, influencing both homologous recombination and nonhomologous end joining (NHEJ) (39). The Mre11 complex's diverse functions in the DNA damage response are likely predicated on its physical association with chromatin. In this regard, one of the least-understood roles of the Mre11 complex in mammals is its association with telomeres.
In mammals, telomeric DNA consists of double-stranded TTAGGG repeats ending in a single-stranded 3′ G overhang, and an array of telomere binding proteins called the shelterin complex that function to prevent telomeres from being recognized as DNA breaks (33). DNA of the overhang invades the double-stranded telomeric repeat sequence to form a t-loop structure (14, 32). The formation of the t-loop requires the telomere protection and remodeling proteins that make up the shelterin complex (7), and these may also contribute to telomere length regulation by preventing telomerase access to chromosomal ends.
Data regarding the role of the Mre11 complex at the telomere have implicated the Mre11 complex in several aspects of telomere maintenance and function. For example, it has been suggested that the Mre11 complex may promote formation of the 3′ telomeric overhang by influencing 5′-to-3′ resection of newly replicated chromosome ends (6). In Saccharomyces cerevisiae, the Mre11 complex recruits the ATM orthologue, Tel1, which is in turn required to recruit telomerase (12, 45). Consequently, Mre11 complex deficiency results in telomere shortening. In mammals, recruitment of telomerase is thought to be regulated primarily by the telomeric protein components TRF1, TPP1, and POT1 (24, 46, 53). However, telomere shortening has also been noted to occur in cell lines from Nijmegen breakage syndrome (NBS) patients in which a hypomorphic Nbs1 allele is expressed, leading to the suggestion that the Mre11 complex may also promote telomerase function in mammals (36). The Mre11 complex associates with telomeres through its interaction with the shelterin component Trf2, apparently in a cell cycle-dependent manner (47, 54). The significance of this physical association is unclear, as genetic depletion of Rad50, a component of the Mre11 complex, does not phenocopy depletion of Trf2 in most respects (1).
To examine the function of the Mre11 complex at mammalian telomeres, we established mouse embryonic fibroblasts (MEFs) derived from a mouse expressing the hypomorphic Mre11ATLD1 allele, crossed to telomerase deficient TertΔ/Δ mice (23, 42), and assessed the rate of telomere shortening. Mre11 complex hypomorphism in MEFs did not affect telomere length, irrespective of telomerase status. In Mre11ATLD1/ATLD1 TertΔ/Δ cells, the fusion of eroded telomeres was reduced compared to TertΔ/Δ cells with telomeres shortened to the same extent, suggesting that the Mre11 complex is involved in the response to critically short telomeres. This interpretation was supported by data obtained using a conditional Trf2 allele to generate acute telomere dysfunction in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells. Collectively the data support a role for the Mre11 complex in the recognition and signaling of dysfunctional telomeres. The character of fusions arising in cre-infected Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/F cells further suggests that the Mre11 complex may influence the processing of chromosome ends following DNA replication en route to t-loop formation.
Primary MEFs were generated from appropriate crosses as previously described (43). For simian virus 40 (SV40) transformation, early-passage MEFs were transfected with the p129-Mertz SV40 plasmid by using Lipofectamine 2000 (Invitrogen) in accordance with manufacturer's instructions. For 3T3 transformation and subsequent culture of both SV40- and 3T3-transformed cells, cells were counted and replated every 3 days in accordance with the 3T3 protocol (44). Cumulative population doubling levels (cPDLs) were calculated using the formula ΔPDL = log(nf/no)/log(2), where no is the initial number of cells and nf is the final number of cells (16).
Metaphases were prepared from cultures treated with colcemid (2 × 10−7 M) for 1 h or less for rapidly proliferating cultures. Cells were then trypsinized and hypotonically swelled (0.075 M KCl) for 7 min at 37°C, fixed, washed in ice-cold methanol-acetic acid (3:1), dropped onto slides, and air dried overnight. Slides were rehydrated in phosphate-buffered saline (PBS), dehydrated through a series of ethanol washes (70%, 90%, and 100%), and air dried. Hybridization mixture (10 mM Tris-Cl, pH 7.2, 70% [vol/vol] formamide, 0.5% [wt/vol] block reagent [Dupont NEN], 1:1,000 TelC-fluorescein isothiocyanate [FITC] peptide nucleic acid probe [Applied Biosystems]) was added to slides and denatured at 80°C for 3 min and then hybridized at room temperature for 2 h in a dark, humid chamber. Slides were washed two times for 15 min (70% formamide, 10 mM Tris-Cl, pH 7.2, 0.1% bovine serum albumin) and then three times for 5 min (100 mM Tris-Cl, pH 7.2, 150 mM NaCl, 0.08% Tween, with DAPI [4′,6-diamidino-2-phenylindole] added to the second wash), dehydrated in ethanol as described above, dried, and mounted. A total of >1,000 metaphase chromosomes from a minimum of 25 spreads were analyzed.
Quantitative FISH (qFISH) was carried out as described in reference 35. Briefly, staining was carried out as for telomere FISH, with the following modifications. After air drying overnight, slides were rehydrated in PBS for 15 min, then fixed in 4% formaldehyde in PBS for 2 min, and washed three times for 5 min in PBS. Slides were treated with 1 mg/ml pepsin in 0.01 N HCl, pH 2.0, at 37°C for 10 min, washed two times for 2 min in PBS, then refixed in 4% formaldehyde in PBS, washed three times for 5 min in PBS, and dehydrated in an ethanol series (70%, 90%, and 100%) for 5 min each. Hybridization, washing, and mounting were done as described for telomere FISH.
For image capture and analysis, a reference slide of 0.2-μm FITC fluorescent beads (Molecular Probes) was used to generate control images for calibration of the microscope (see reference 35 for details). Image capture and analysis were carried out using an Axiovert 300 M (Zeiss) microscope and Volocity software (Improvision). Captured images did not contain saturated pixels. Telomere fluorescence units (TFU) were measured using Volocity software. Telomere regions to be measured were manually assigned and intensity was measured by Volocity to obtain the average number of TFU for each telomeric region. Data from all slides were pooled to give telomere length profiles for the cell population.
Chromosome orientation-FISH (CO-FISH) was done as described for telomere FISH above, with the following modifications. Infected cells were grown in Dulbecco's modified Eagle's medium (DMEM) plus 10% cosmic calf serum (CCS) containing a 3:1 ratio of 5′-bromo-2′-deoxyuridine (BrdU)-5′-bromo-2′-deoxycytidine (BrdC) (Sigma) at a final concentration of 1 × 10−5 M for 17 h, with colcemid (2 × 10−7 M) added for the last hour. Metaphase spreads were prepared as described above, except that care was taken not to denature the DNA after the chromosomes were dropped. Slides were rehydrated in PBS, then treated with 0.5 mg/ml RNase A in PBS at 37°C for 10 min, stained with 0.5 μg/ml Hoechst 33258 (Sigma) in 2× standard saline citrate (SSC) (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate) for 15 min at room temperature, and exposed to 365 nm UV light (Stratalinker 1800 UV irradiator) in 2× SSC for 30 min. The BrdU/BrdC-substituted DNA was digested with exonuclease III (10 U/μl; Promega) in buffer supplied by manufacturer (50 mM Tris-HCl, pH 8.0, 5 mM MgCl2, and 5 mM dithiothreitol) for 10 min at room temperature. Slides were then rinsed in PBS and dehydrated through an ethanol series as before. Hybridization mixture (10 mM Tris-Cl, pH 7.2, 70% [vol/vol] formamide 0.5% [wt/vol] block reagent [Dupont NEN], 1:10,000 TelG-Cy3 peptide nucleic acid probe [Applied Biosystems]) was added, and TelG probe was hybridized at room temperature for 2 h in a dark, humid chamber. Slides were quickly rinsed in wash I (70% [vol/vol] formamide, 10 mM Tris-Cl, pH 7.2, 0.1% [wt/vol] bovine serum albumin), and then hybridization mixture with 1:1,000 TelC-FITC probe was applied for 2 h in a dark, humid chamber. Slides were then washed twice with wash I for 30 min each and three times with wash II for 5 min each at room temperature, with DAPI added to the second wash. Slides were dehydrated and mounted as described above.
Clamped homogenous electric field gel electrophoresis of mouse DNA was performed essentially as described previously (48). Cells were resuspended in PBS and mixed 1:1 (vol/vol) with 2% (wt/vol) pulsed-field gel grade agarose (Bio-Rad) to obtain 1 × 106 cells per agarose plug. Plugs were digested with 1 mg/ml proteinase K in TENS buffer (100 mM EDTA, 0.2% [wt/vol] sodium deoxycholate, 1% [vol/vol] sodium lauryl sarcosine) and washed extensively in TE buffer (10 mM Tris, pH 8, 1 mM EDTA). Plugs were incubated overnight at 37°C with 60 U MboI in 0.5 ml buffer. Following digestion, plugs were washed in TE buffer and equilibrated in 0.5× Tris-borate-EDTA (TBE) before being loaded into a 1% [wt/vol] agarose-0.5× TBE gel. The gel was run using a CHEF-DRII pulsed-field gel electrophoresis apparatus (Bio-Rad) in 0.5× TBE for 18 h with the following settings: initial pulse, 5 s; final pulse, 5 s; 5.4 V/cm2; and 14°C. In-gel hybridization with a [γ-32P]ATP end-labeled (CCCTAA)4 oligonucleotide and subsequent denaturation and hybridization steps were performed as described previously (18). Gels were exposed onto a PhosphorImager screen, and lanes were quantified with ImageGauge software.
Lentiviral production, concentration, and determination of titers were carried out using established methods (10, 25). SV40-transformed MEFs were infected with either a cre-green fluorescent protein vector-based lentivirus or the same virus with a deleted cre locus as a negative control. Cells were infected in suspension at 1 × 106 cells/ml at a multiplicity of infection of 10 in DMEM plus 10% CCS with 5 μg/ml polybrene. Tubes were spun at 1,900 rpm (~600 × g) for 90 min, with occasional stops to manually resuspend cells. After viral infection, cells were plated and grown in DMEM containing 10% CCS.
Protein lysates for Rap1 were prepared by subjecting cells to three freeze-thaw cycles in high-salt-concentration TNG buffer (50 mM Tris, pH 7.5, 400 mM NaCl, 0.1% Tween, and 0.5% NP-40 plus protease inhibitors) and transferred onto a nitrocellulose membrane. To prepare extracts for γ-H2AX detection, the chromatin pellet from regular lysates was further incubated for 30 min in 0.1 M HCl, and the resulting supernatant was used for immunoblotting. Binding and washing steps were done with 5% milk and PBS-0.05% Tween 20 buffer. Rabbit anti-Rap1 polyclonal (1:10,000; obtained from T. de Lange), antiactin mouse monoclonal (1:1,000; AC-40; Sigma), anti-γ-H2AX mouse monoclonal (1:500; Ser139; Upstate), and anti-total histone H2A rabbit (1:500; Upstate) primary antibodies and horseradish peroxidase-conjugated species-specific secondary antibodies (Pierce) were used. Horseradish peroxidase signal was detected with ECL Plus reagent (Amersham).
Cells were grown on coverslips, fixed for 15 min in 4% paraformaldehyde, washed in PBS, and blocked for 30 min in blocking solution (1 mg/ml bovine serum albumin, 3% goat serum, 0.1% Triton X-100, 1 mM EDTA). Coverslips were incubated at room temperature for 1 h with polyclonal anti-53BP1 rabbit antibody (1:1,000 diluted in blocking solution; Novus), washed, incubated with anti-rabbit secondary antibody (Rhodamine, diluted in blocking solution 1:200) for 30 min, fixed for 5 min in 4% paraformaldehyde, and washed in PBS. Coverslips were dehydrated through a series of ethanol washes (70%, 90%, and 100%), hybridized and denatured for 5 min at 80°C, and then hybridized for 2 h at room temperature by using a FITC-TelC probe (1:1,000; Applied Biosystems). Coverslips were washed twice each for 15 min in FISH wash solution (70% formamide, 10 mM Tris-Cl, pH 7.2) and then three times in PBS, with DAPI added to the second wash, and were dried and mounted.
Spectral karyotyping (SKY) of metaphase chromosomes was carried out as described previously (43).
Statistical significance was determined by Fisher's exact test (for 3T3 fusions, cre-induced fusions, and telomere dysfunction-induced focus [TIF] formation), the Wilcoxon rank sum test (for qFISH), and the chi-square goodness-of-fit test (for orientation of fusions), using Excel or Mstat software (Norman Drinkwater, McArdle Laboratory for Cancer Research).
Cells from the hypomorphic mouse mutant Mre11ATLD1/ATLD1 exhibit defects in the ATM-dependent DNA damage response (29, 39, 42). Cells from this mutant, as well as an additional Mre11 complex hypomorph, the Nbs1ΔB/ΔB mouse, which is also defective in ATM activity (49), were used to examine the telomeric functions of the Mre11 complex in mammals. Mre11+/ATLD1 mice were crossed with telomerase heterozygous (Tert+/Δ) mice to create double heterozygous mutant mice, and these animals were then crossed together to generate Mre11ATLD1/ATLD1 Tert+/+, Mre11+/+ TertΔ/Δ, and Mre11ATLD1/ATLD1 TertΔ/Δ MEFs.
MEFs were isolated and passaged by following the 3T3 protocol (44), and cPDLs were calculated for each genotype throughout the course of the experiment. Individual embryo cultures were transformed both by SV40 infection and by classic 3T3 transformation. Although 3T3 transformation was slower than SV40 transformation (approximately passage 20 for 3T3 cells versus approximately passage 7 for SV40 cells) (Fig. (Fig.1A),1A), no overt differences in the efficiency of transformation of any genotype or in the growth rate of the cells in culture were observed (Fig. (Fig.1A).1A). To ensure that meaningful cPDL comparisons could be made, the rates of BrdU incorporation and cell death were determined throughout the course of the experiment by flow cytometry. For 3T3-transformed cultures, 49% of cells were BrdU positive on average throughout the experiment. For SV40-transformed cultures, an average of 56% of cells were BrdU positive (Table (Table1).1). For both transformation methods, cells with sub-G1 DNA content (correlated with cell death) were generally less than 2% of the culture (Table (Table1).1). These parameters confirm that the potentially confounding effects of differing cell death rates among the various genotypes did not skew the calculated cPDLs and that the calculated values reflected comparable numbers of cell divisions among the genotypes examined.
It has previously been reported that wild-type (WT) (telomerase-proficient) MEFs display minimal telomere shortening upon continual passage in tissue culture (4). To determine the effect of Mre11 complex hypomorphism, metaphase spreads were analyzed following hybridization of a FITC-labeled telomeric probe (TelC) at cPDLs 70, 300, and 400 and were assessed for the appearance of signal-free ends and telomere fusions. At cPDL 70, none of the genotypes displayed marked loss of telomere signal or telomere fusion formation. At late passage (cPDL 300), Mre11ATLD1/ATLD1 cells did not show loss of telomere signal, nor did we observe the appearance of chromosome fusions (Fig. 1B and D).
As expected, the telomeres of TertΔ/Δ and Mre11ATLD1/ATLD1 TertΔ/Δ MEFs underwent progressive shortening upon passage through culture (4, 23), resulting in the appearance of signal-free ends and chromosomal fusions at high cPDLs (Fig. 1B and D). Telomere shortening in cPDL 70 and 300 MEFs was quantified by qFISH (19). We did not observe telomere shortening in Mre11ATLD1/ATLD1 MEFs at cPDL 300 (Fig. (Fig.1C,1C, top row), indicating that Mre11 complex hypomorphism does not appear to impair the recruitment of telomerase. In TertΔ/Δ MEFs, marked telomere shortening was observed, from an average TFU count of 766 at cPDL 70 to an average TFU count of 520 TFU at cPDL 300. Mre11ATLD1/ATLD1 TertΔ/Δ cultures also showed significant telomere shortening at cPDL 300, and there was no difference in telomere length at this cPDL between Mre11+/+ TertΔ/Δ (520 TFU) and Mre11ATLD1/ATLD1 TertΔ/Δ (517 TFU) MEFs (Fig. (Fig.1C,1C, bottom two rows).
These results were confirmed using TRF Southern blotting, which showed that equivalent levels of shortening were observed between Mre11+/+ TertΔ/Δ and Mre11ATLD1/ATLD1 TertΔ/Δ MEFs (data not shown). TRF analysis also confirmed that maintenance of the G-strand overhang was unaffected (data not shown). These data support the interpretation that the murine Mre11 complex does not influence telomere length maintenance in either telomerase-proficient or telomerase-deficient mouse cells.
As described above, Mre11+/+ TertΔ/Δ and Mre11ATLD1/ATLD1 TertΔ/Δ MEFs showed diminished telomere signals due to telomere shortening and the appearance of chromosomal fusions (Fig. 1B and D). The observed fusions did not contain cytologically detectable telomeric sequences, suggesting that they result from telomere attrition-dependent uncapping. Fusions were predominantly p-p fusions (short arm) (Fig. (Fig.1E),1E), which may be a reflection of the increased stability of p-p fusions in mouse cells in culture. Short arm fusions are likely to be more stable because of the acrocentric nature of mouse telomeres; thus, p-q and q-q fusions create dicentric telomeres that are more likely to undergo breakage upon cell division (27). Analysis by SKY of cPDL 300 spreads demonstrated that the observed fusions were not clonal, as we did not detect rearrangements common to multiple cells (Table (Table2).2). This suggests that fusion of critically short telomeres is ongoing in late passage cells.
The rate of telomere fusion was reduced in Mre11ATLD1/ATLD1 TertΔ/Δ cells. At cPDL 300, 5% of all chromosomes were fused in Mre11ATLD1/ATLD1 TertΔ/Δ, in comparison to 8% of all chromosomes in TertΔ/Δ (P < 0.001). Fewer fusions were also observed in cPDL 400 Mre11ATLD1/ATLD1 TertΔ/Δ than in cPDL 400 Mre11+/+ TertΔ/Δ (P < 0.0003) (Fig. (Fig.1D).1D). These data suggest a role for the Mre11 complex in NHEJ of dysfunctional telomeres.
The Mre11 complex associates with mammalian telomeres and has been suggested to recognize uncapped telomeres as DNA damage following replication (47, 54), resonant with its role in the recognition of interstitial DSBs and the subsequent activation of ATM (21, 39). It has recently been shown that loss of the shelterin component Trf2 in the absence of ATM results in a significant decrease in telomere fusion formation (8). On these bases, we hypothesized that the Mre11 complex is required for recognition of dysfunctional telomeres and activation of ATM; hence, the observed decrease in telomere fusions in late-passage Mre11ATLD1/ATLD1 TertΔ/Δ MEFs would reflect the decrement in ATM activity attendant to the Mre11ATLD1 allele.
To test this hypothesis, we utilized the Trf2 conditional allele, Trf2Flox, which provides an alternative means of generating telomere dysfunction, and which may elicit a response distinct from that seen at eroded telomeres. Introduction of lentiviral cre recombinase into Trf2F/Δ or Trf2F/F cells (which were used interchangeably here) generates Trf2-null cells (Trf2Δ/Δ) and results in acute telomere dysfunction due to loss of Trf2 and subsequent telomere uncapping (5). The uncapped telomere activates the DNA damage response and leads to chromosome fusions and the formation of TIF (41). We generated Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/F MEFs and assessed their responses to acute telomere dysfunction.
Trf2F/F cells were infected with lentivirus expressing the cre recombinase, and depletion of Trf2 protein was monitored by assessing the level of Rap1. Mouse Trf2 antisera recognize a nonspecific band that obscures Trf2 on Western blots. As Rap1 is destabilized upon loss of Trf2, Rap1 is used as a surrogate marker for Trf2 depletion (5) (Fig. (Fig.2A).2A). As previously shown, depletion of Trf2 in WT cells results in the formation of multiple chromosome fusions with retention of telomere signal at the fusion junction (Fig. (Fig.2B)2B) (20). In this study, 43.3% of the total chromosomes were fused in Trf2Δ/Δ cells (chromosome plus chromatid type fusions) (Fig. (Fig.2D).2D). Consistent with the reduction in fusions seen in cells with critically short telomeres, both Mre11ATLD1/ATLD1 Trf2Δ/Δ and Nbs1ΔB/ΔB Trf2Δ/Δ MEFs exhibited significant reductions in total fusions, to 18.3% (P < 0.0003 for comparison with Trf2Δ/Δ) and 7.0% (P < 0.0001 for comparison with Trf2Δ/Δ), respectively (Fig. (Fig.2D2D).
Assessment of telomere fusions was also carried out using TRF Southern blots. TRF length analysis showed increased fusion formation in Trf2Δ/Δ cells, as evidenced by the higher-molecular-weight smear indicated on the blot and the loss of the WT telomere bulk. In contrast, Mre11ATLD1/ATLD1 Trf2Δ/Δ and Nbs1ΔB/ΔB Trf2Δ/Δ MEFs showed reduced fusion formation (Fig. (Fig.2C,2C, right panel, lanes 10 and 12), consistent with the data obtained from metaphase cells by using telomere FISH. The single-stranded G overhang (Fig. (Fig.2C,2C, left panel, lanes 4 and 6) remained intact in both cre-infected Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/F cells, showing that 3′ overhang loss does not occur upon acute telomere dysfunction in either Mre11ATLD1/ATLD1 Trf2Δ/Δ or Nbs1ΔB/ΔB Trf2Δ/Δ cells.
Although overall fusions were markedly reduced in cre-infected Mre11ATLD1/ATLD1 Trf2F/Δ or Nbs1ΔB/ΔB Trf2F/F, a substantial fraction of the residual fusions in those mutants were events in which only one chromatid is fused. Less than 2.0% of the total fusions in Trf2Δ/Δ were chromatid fusions, whereas 48.5% of Mre11ATLD1/ATLD1 Trf2Δ/Δ (P < 0.0001 for comparison with Trf2Δ/Δ) and 29.3% of Nbs1ΔB/ΔB Trf2Δ/Δ (P < 0.0001 for comparison with Trf2Δ/Δ) fusions were of this class (Fig. (Fig.2B2B).
To examine the basis of the observed bias toward chromatid type fusions, these events were characterized using CO-FISH, which allows for identification of the leading (5′) and lagging (3′) strands (2). If fusion formation were a random event, a 1:2:1 ratio of leading-leading, leading-lagging, and lagging-lagging fusions would be observed (Fig. (Fig.3A3A).
The rare chromatid type fusions seen in Trf2Δ/Δ cells (fewer than one per spread) were equally distributed among leading-leading, leading-lagging, and lagging-lagging chromatid fusions (Fig. 3A and B). It is likely that this departure from expectation is an artifact of the small sample size available. In contrast, Mre11ATLD1/ATLD1 Trf2Δ/Δ and Nbs1ΔB/ΔB Trf2Δ/Δ MEFs exhibited a striking tendency toward leading-leading-strand fusions (Fig. 3B and C and Table Table3).3). In Mre11ATLD1/ATLD1 Trf2Δ/Δ MEFs, 96.1% of chromatid fusions and 87.3% of Nbs1ΔB/ΔB Trf2Δ/Δ MEFs were leading-leading-strand fusions (P < 0.0001 for comparison with Trf2Δ/Δ). These data indicate that fusions involving the lagging strand are strongly inhibited in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells.
Telomeres protect chromosome ends from being recognized as DNA DSBs. Accordingly, telomere dysfunction in mammalian cells induces a DNA damage response that can be measured through the formation of TIF, consisting of telomeric 53BP1 or γ-H2AX foci (5). Given the role of the Mre11 complex in promoting both the activation and the activity of ATM, we asked whether the Mre11 complex is required for the response to dysfunctional telomeres. TIF formation in cre-infected Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/F cells was examined, and as previously seen in Atm−/− cells, both showed drastically reduced TIF formation (Fig. (Fig.4A).4A). As expected, 60.0% of Trf2Δ/Δ MEFs were TIF positive (defined as cells having at least five telomeric 53BP1 foci) following Trf2 deletion. In contrast, only 16.8% and 24.0% of cells were TIF positive in Mre11ATLD1/ATLD1 Trf2Δ/Δ and Nbs1ΔB/ΔB Trf2Δ/Δ, respectively (P < 0.0001 for comparison with Trf2Δ/Δ) (Fig. (Fig.4B).4B). The formation of γ-H2AX induced by Trf2 depletion was similarly impaired in Mre11 complex mutants (Fig. (Fig.4C).4C). These observations suggest that the Mre11 complex's DNA damage recognition and signaling functions, required for the response to interstitial DNA damage, are also required for the detection or signaling of acute telomere dysfunction. Moreover, the data underscore the intimate relationship of the DNA damage response pathway and telomere function.
In this study, we employed two experimental systems to examine the functions of the Mre11 complex at mammalian telomeres. We present evidence that the Mre11 complex is required for the response to telomere dysfunction and further that the disposition of telomeric ends following DNA replication may be influenced by the Mre11 complex. First, the effect of Mre11 complex hypomorphism was examined during long-term serial passage of telomerase-proficient and -deficient (TertΔ/Δ) cells. No telomere attrition was seen in immortalized Mre11ATLD1/ATLD1 cells over the course of over 400 population doublings, suggesting that the Mre11 complex does not influence telomere length regulation in murine cells. This is in apparent contrast to data from human cells showing that Nbs1, a component of the Mre11 complex, promotes telomerase-mediated telomere synthesis and, when defective, leads to shortened telomeres (36). Similarly, the degree of telomere attrition in Mre11ATLD1/ATLD1 TertΔ/Δ was indistinguishable from that in TertΔ/Δ alone, further supporting the view that the Mre11 complex does not contribute to telomere length maintenance.
Whereas Mre11 complex hypomorphism did not overtly affect normal telomeres, several lines of evidence indicated that the complex governs the cellular response to dysfunctional telomeres. First, propagation of telomerase-deficient cells to the point of telomere dysfunction led to telomere fusions that were at least partially dependent on Mre11, as telomere fusions were less frequent in late-passage Mre11ATLD1/ATLD1 TertΔ/Δ cells than in TertΔ/Δ cells. This indicates that in WT cells, the Mre11 complex recognizes and promotes the “repair” (i.e., fusion) of dysfunctional telomeres. The reduction in fusions was not attributable to differences in telomere length or the disposition of the 3′ single-stranded DNA overhang. The same effect was observed, and was significantly more pronounced, upon the induction of acute telomere dysfunction in Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/F cells, indicating that the Mre11 complex responds to both acute and nonacute telomere dysfunction.
The inhibition of telomere fusions seen here is reminiscent of the effect of Mre11 complex hypomorphism on chromosome fusions associated with DNA-protein kinase catalytic subunit deficiency. The rate of chromosome fusions is elevated in immortalized DNA-protein kinase catalytic subunit-deficient cells (3, 11, 13), and this effect was significantly reduced in Nbs1ΔB/ΔB Prkdcscid/scid cells (40). We propose that the reduction in telomere fusions observed here suggests a role for the Mre11 complex in NHEJ, consistent with recent studies of lymphoid cells from Nbs1ΔB/ΔB and Mre11ATLD1/ATLD1 mice (9, 17, 40). Because they retain extensive telomere sequence, acutely dysfunctional telomeres are more likely to fuse via the canonical DNA ligase IV-dependent pathway. Accordingly, chromatid fusions in cells depleted of both Trf2 and Nbs1 are DNA ligase IV dependent; hence, the same is likely to be true for chromatid fusions in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB. Given that the fusion of both acutely dysfunctional and eroded telomeres is inhibited by Mre11 complex hypomorphism, the most reasonable interpretation is that the underlying mechanisms of fusion are the same in both contexts. Nevertheless, we cannot rule out the formal possibility that Mre11 complex hypomorphism has similar effects on two distinct mechanisms of NHEJ at chromosome ends.
Given that telomere uncapping elicits a DNA damage checkpoint response (33), we propose that the checkpoint defects associated with Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB mutations may at least partially account for the preponderance of chromatid fusions among the residual joining events observed. The G1/S checkpoint is inactive in all cells as a result of SV40 immortalization. Hence, WT G1 cells with uncapped telomeres will undergo fusions prior to S phase or enter S phase with uncapped telomeres. The latter will likely activate the intra-S and/or the G2/M checkpoint. The detection of TIF in interphase cells supports this scenario. Subsequent fusion of those uncapped telomeres would relax the checkpoint and allow cells to progress to metaphase with both sisters fused (i.e., with chromosome fusions). Concurrently, cells with persistent uncapped telomeres will accumulate at the G2/M boundary and simply not progress to metaphase (Fig. (Fig.5A).5A). The intra-S and G2/M checkpoints are both defective in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells (42, 49). Therefore, the progression of Mre11 complex mutant cells entering S phase with uncapped telomeres will not be impeded to the same extent as WT cells, and so metaphase spreads with chromatid fusions are more likely to be obtained (Fig. (Fig.5A).5A). Consistent with this view, the intra-S and G2/M checkpoint defects are more severely impaired in Mre11ATLD1/ATLD1 than in Nbs1ΔB/ΔB (38), and the yield of chromatid fusions is greater (48.5% versus 29.3%) (Fig. (Fig.2B)2B) in those cells.
Although the significant increase in chromatid fusions seen in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells is likely to reflect checkpoint defects in those cells, it does not account for the strong bias toward fusion of leading-strand telomeres. We propose that the structure of the newly replicated telomeric ends in Mre11 complex mutants may influence this outcome. Replication of the leading-strand telomere presumably results in a blunt end, while semicontinuous lagging-strand replication creates a 3′ single-stranded overhang as a result of the terminal Okazaki fragment. This difference in the leading and lagging termini is transient, as resection of both ends to create the 3′ G strand overhang occurs (26).
The bias for leading-strand fusions is consistent with two possibilities (Fig. (Fig.5B).5B). First, resection of the leading-strand telomere to create the 3′ overhang may be delayed in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells. The presence of the 3′ overhang is likely to impair NHEJ-mediated fusion; hence, delayed resection of the blunt end would favor NHEJ of the leading-strand telomere. The alternative, and nonexclusive, possibility is that lagging-strand fusions are inhibited in Mre11 complex mutants because the complex also promotes degradation the 3′ overhang of unprotected telomeres and thereby promotes NHEJ of lagging-strand telomeres. The predominance of chromosome fusions in all genotypes upon Trf2 deletion (Fig. (Fig.2D)2D) suggests that ≥50% of 3′ overhangs are degraded to permit fusion. This could reflect the hypomorphic nature of the Mre11 complex mutants examined as well as redundancy with other nucleases.
It is unlikely that the Mre11 complex per se mediates telomeric resection. It is clear that bulk resection of DNA ends in S. cerevisiae is not mediated by the Mre11 complex (22, 30, 31), that the polarity of Mre11-dependent exonuclease activity is 3′ to 5′ but that 5′-to-3′ polarity is required (34), and that telomeric end resection is unaffected in mre11Δ yeast strains (15). Nevertheless, in conjunction with Sae2 (or CtIP in mammals), the Mre11 nuclease appears to initiate the resection process via endonucleolytic removal of a terminal oligonucleotide prior to bulk resection by Exo1, Dna2, and the helicase Sgs1 (28, 37, 55). This initiation step may be delayed in Mre11 complex mutants, or recruitment of the required activities to the telomere may be impaired in Mre11ATLD1/ATLD1 and Nbs1ΔB/ΔB cells. In either scenario, the data support the view that the Mre11 complex influences resection of the leading-strand telomere. Whether this apparent effect on telomeric end processing can fully account for the overall reduction in fusions is not clear. In this regard, the complex's role in the bridging of DNA ends is also likely to at least partially underlie these effects (50-52).
Finally, the data presented herein indicate that the Mre11 complex is required for activation of the response to dysfunctional telomeres. TIF formation in cre-infected Mre11ATLD1/ATLD1 Trf2F/Δ and Nbs1ΔB/ΔB Trf2F/Δ cells was strongly impaired in response to Trf2 deletion. This result phenocopies the outcome seen in cre-infected Atm−/− Trf2F/Δ cells (8). Whereas many aspects of Atm-dependent responses to telomere uncapping, such as the regulation of NHEJ, are specific to the telomere (8), the observation that the Mre11 complex is required for the response to telomere dysfunction suggests that the early events in this response overlap significantly with those during the response to interstitial DNA damage.
This work was supported by NIH grants to J.H.J.P. C.L.A. is a Leukemia & Lymphoma Society Special Fellow.
We thank Saurav De for animal care and insights; Lea Harrington and Natalie Erdmann for TertΔ/Δ mice and qFISH analysis (NIH R01 AG02398); Titia de Lange for Trf2Flox mice, data, and discussions prior to publication; Margaret Leversha for SKY analysis; Eros Lazzerini Denchi for technical advice; and members of our laboratory for insightful discussion.
Published ahead of print on 10 August 2009.