Exonuclease-1 Deletion Impairs DNA Damage Signaling and Prolongs Lifespan of Telomere-Dysfunctional Mice

Exonuclease-1 Deletion Impairs DNA Damage Signaling and Prolongs Lifespan of Telomere-Dysfunctional Mice Sonja Schaetzlein,1,7 N.R. Kodandaramireddy,1,7 Zhenyu Ju,1,8 Andre Lechel,1 Anna Stepczynska,1 Dana R. Lilli,3 Alan B. Clark,4 Cornelia Rudolph,2 Florian Kuhnel,1 Kaichun Wei,5 Brigitte Schlegelberger,2 Peter Schirmacher,6 Thomas A. Kunkel,4 Roger A. Greenberg,3 Winfried Edelmann,5 and K. Lenhard Rudolph1,* 1Department

of Gastroenterology, Hepatology and Endocrinology of Cell and Molecular Pathology Medical School Hannover, 30625 Hannover, Germany 3Department of Cancer Biology, Abramson Family Cancer Research Institute, University of Pennsylvania School of Medicine, Philadelphia, PA 19104-6160, USA 4Laboratory of Structural Biology, National Institute of Environmental Health Sciences, Research Triangle Park, NC 27709, USA 5Department of Cell Biology, Albert Einstein College of Medicine, Bronx, NY 10461, USA 6Institute of Pathology, University Hospital, Im Neuenheimer Feld 220/221, 69120 Heidelberg, Germany 7These two authors contributed equally to this work. 8Present address: Institute of Laboratory Animal Science, Chinese Academy of Medical Science, Chaoyang District, 100021 Beijing, China. *Correspondence: [email protected] DOI 10.1016/j.cell.2007.08.029 2Institute

SUMMARY

Exonuclease-1 (EXO1) mediates checkpoint induction in response to telomere dysfunction in yeast, but it is unknown whether EXO1 has similar functions in mammalian cells. Here we show that deletion of the nuclease domain of Exo1 reduces accumulation of DNA damage and DNA damage signal induction in telomeredysfunctional mice. Exo1 deletion improved organ maintenance and lifespan of telomeredysfunctional mice but did not increase chromosomal instability or cancer formation. Deletion of Exo1 also ameliorated the induction of DNA damage checkpoints in response to g-irradiation and conferred cellular resistance to 6-thioguanine-induced DNA damage. Exo1 deletion impaired upstream induction of DNA damage responses by reducing ssDNA formation and the recruitment of Replication Protein A (RPA) and ATR at DNA breaks. Together, these studies provide evidence that EXO1 contributes to DNA damage signal induction in mammalian cells, and deletion of Exo1 can prolong survival in the context of telomere dysfunction. INTRODUCTION Telomeres form the ends of eukaryotic chromosomes (Blackburn, 2001). The main function of telomeres is to cap chromosomal ends, thus preventing induction of

DNA damage pathways. Telomere capping function depends on telomere length and telomere structure (de Lange, 2005). Due to the end replication problem of DNA polymerase and the processing of telomeres during cell cycle progression, telomeres shorten with each round of cell division (Shay and Wright, 2000). Telomere shortening limits the proliferative lifespan of primary human cells (Allsopp et al., 1992) and can reduce regenerative reserve and organ homeostasis during aging and chronic diseases (for review see Djojosubroto et al., 2003). In mammalian cells telomere shortening leads to telomere dysfunction inducing senescence or apoptosis (Lechel et al., 2005; Lee et al., 1998; Wright and Shay, 1992). Critically short telomeres lose chromosomecapping function. The most upstream response to telomere uncapping is the formation of DNA damage foci at dysfunctional telomeres (d’Adda di Fagagna et al., 2003; Takai et al., 2003). These DNA damage foci activate downstream DNA damage signals, which involve the ATM and ATR kinases (d’Adda di Fagagna et al., 2003) inducing the p53 pathway (Chin et al., 1999; Vaziri and Benchimol, 1996). Cell cycle arrest and apoptosis in response to telomere dysfunction represent a tumor suppressor mechanism (Wright and Shay, 1992). As a downside, these checkpoints may contribute to decreased regenerative reserve and impaired organ maintenance in response to telomere dysfunction and aging (Choudhury et al., 2007). The generation of telomerase-deficient mice, which carry a homozygous deletion of the telomerase RNA component (mTerc/), has delivered a unique tool to study consequences of telomere dysfunction in vivo. Late generation (G3) mTerc/ mice exhibit cellular and molecular phenotypes induced by telomere dysfunction, including

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 863

activation of the p53-DNA-damage pathway, impaired proliferation, increased apoptosis, and chromosomal fusions (Lee et al., 1998; Rudolph et al., 1999; Chin et al. 1999). Mice with dysfunctional telomeres show impaired maintenance of organs with high rates of cell turnover and premature aging of these compartments (Lee et al., 1998; Rudolph et al., 1999; Choudhury et al. 2007; Hao et al. 2005; Hemann et al. 2001). Telomere-dysfunctional mice have been used to analyze the function of different components of the DNA damage pathway in the context of telomere dysfunction in vivo (Qi et al., 2003; Wong et al., 2003; Chin et al., 1999; Artandi et al., 2000; Greenberg et al., 1999; Choudhury et al., 2007). These studies have shown that activation of the p53 pathway has a major role in mediating the adverse effects of telomere dysfunction on organismal aging, but at the same time, this pathway has a pivotal role in preventing the evolution of chromosomal instability and cancer initiation in response to telomere dysfunction (Chin et al., 1999; Artandi et al., 2000). Of note, ATM deletion did not rescue premature aging of telomere-dysfunctional mice (Wong et al. 2003), indicating that ATM-independent pathways can activate p53 in response to telomere dysfunction possibly involving ATR (d’Adda di Fagagna et al., 2003) —a gene involved in activation of DNA damage signals in response to generation of single-stranded DNA (ssDNA) (Zou et al., 2003). The induction of chromosomal instability and tumor initiation in response to telomere dysfunction is linked to the evolution of chromosomal instability and the formation of nonreciprocal translocation (Artandi et al., 2000). This response appears to be initiated by DNA repair pathways that induce chromosomal fusions in response to telomere uncapping, with the consequence of fusion-bridge-breakage cycles (Maser et al., 2007a; Smogorzewska et al., 2002). Exonuclease-1 (EXO1) is a 50 -30 exonuclease. A role of EXO1 for the induction of checkpoints in response to telomere dysfunction has been revealed in studies on yeast. Deletion of Exo1 rescued the induction of senescence in telomere-dysfunctional yeast strains (Maringele and Lydall, 2002). In addition, EXO1 processing of dysfunctional telomeres has been implicated in generating chromosomal translocation in response to telomere dysfunction in yeast (Hackett and Greider, 2003). EXO1 can also mediate survival of telomere-dysfunctional yeast strains by increasing homologous recombination at dysfunctional telomeres (Maringele and Lydall, 2004). Knockout mice carrying a deletion mutation of the nuclease domain of Exo1 are viable, but show defects in DNA mismatch repair, accompanied by slightly increased cancer susceptibility at advanced age (Wei et al., 2003). The role of EXO1 in response to telomere dysfunction and DNA breaks in mammalian cells is unknown. Here we show that Exo1 deletion rescues the accumulation of DNA damage in telomere-dysfunctional mice, as well as DNA damage signal induction in response to g-irradiation (g-IR) and at laser-induced DNA breaks. In vivo, Exo1 deletion improved organ maintenance and prolonged life-

864 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

span of telomere-dysfunctional mice without accelerating chromosomal instability or cancer formation. RESULTS Exo1 Deletion Prolongs the Lifespan of Mice with Dysfunctional Telomeres without Accelerating Tumor Formation Mice carrying a deletion of the nuclease domain of Exo1 (DExon 6 = Exo1+/, Wei et al., 2003) were crossed for three generations with mTerc/ mice on a C57BL/6J background. As previously reported the lifespan of G3 mTerc/ mice (n = 25) was shortened compared with mTerc+/+, Exo1+/+ mice (n = 89, p < 0.0001, Figure 1A). Homozygous Exo1 deletion (Exo1/) shortened the lifespan of mTerc+/+ mice (p = 0.009, Figure 1A; Wei et al., 2003). An increase in spontaneous cancer has been reported in Exo1/ mice of a mixed genetic background (Wei et al. 2003). In agreement with the relative cancer resistance of the C57BL/6J mouse strain, the overall cancer incidence was low in our mouse cohorts (Figure 1B and Table S1 in the Supplemental Data, available with this article online), and Exo1 deletion did not result in a significant increase in cancer formation, indicating that other factors may contribute to the lifespan reduction in mTerc+/+, Exo1/ mice. In contrast to the effects on lifespan of mTerc+/+ mice, deletion of Exo1 significantly prolonged survival of G3 mTerc/, Exo1/ mice (n = 34) compared with G3 mTerc/, Exo1+/+ mice (p < 0.0001, Figure 1A). This rescue in lifespan was associated with an improvement of overall fitness, as reflected by an increase in body weight of aging G3 mTerc/, Exo1/ mice compared with G3 mTerc/, Exo1+/+ mice (p = 0.0002, Figure S1A in the Supplemental Data available with this article online). Notably, Exo1 deletion did not accelerate tumor formation in G3 mTerc/ mice (Figure 1B, p = 0.3). Exo1 Deletion Rescues Organ Maintenance in Aging Telomere-Dysfunctional Mice G3 mTerc/ mice are phenotypically normal at the age of 3 months, but show an atrophy of the intestinal epithelium and impaired hematolymphopoiesis at the age of 12–15 months (Rudolph et al., 1999; Choudhury et al. 2007). The evolution of these phenotypic changes correlates with the progression of telomere dysfunction in aging G3 mTerc/ mice. A survey of our aging cohorts showed that Exo1 deletion rescued organ maintenance in 12- to 15-month-old G3 mTerc/ mice (Figures 1C–1F and Figures 2A–2F). In a cohort of 24-month-old G3 mTerc/, Exo1/ mice (n = 4), not in 24-month-old mTerc+/+ mice (Figures 1D and 1F and Figures 2B and 2C), some of the phenotypes of impaired organ maintenance appeared, indicating that Exo1 deletion transiently rescued organ maintenance in G3 mTerc/ mice, but that EXO1-independent mechanisms limited organ maintenance in very old G3 mTerc/, Exo1/ mice. Deletion of Exo1 did not rescue testis atrophy in G3 mTerc/ mice because Exo1 deletion itself causes

Figure 1. Exo1 Deletion Prolongs the Lifespan of Telomere-Dysfunctional Mice without Accelerating Cancer Formation (A) Survival curves for mTerc+/+, Exo1+/+ (n = 89), mTerc+/+, Exo1/ (n = 27), G3, Exo1+/+ (n = 25), and G3, Exo1/ mice (n = 34). (B) Tumor-free survival curves for the indicated mouse cohorts. (C) Representative photographs of whole-mount staining of the colon of 12- to 15-month-old mice of the indicated genotypes (magnification bar, 500 mm). (D) Histogram of the number of crypts per low-power vision field (353) in the colon of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–6 mice per group; data are shown as mean; error bars represent standard deviation [SD]). (E) Representative photographs of H&E-stained longitudinal sections of the small intestine of 12- to 15-month-old mice of the indicated genotypes (magnification bar, 200 mm). (F) Histogram of the number of basal crypts per vision field (1003) in the small intestine of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD).

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 865

Figure 2. Exo1 Deletion Rescues Lymophopoiesis in Aging Telomere-Dysfunctional Mice (A) Representative photographs of H&E-stained longitudinal sections of the spleen of 12- to 15-month-old mice of the indicated genotypes (magnification bar, 500 mm). The lymphocytes containing white pulp are circled. (B) Histogram of the frequency of B lymphocytes in spleen of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–6 mice per group; data are shown as mean; error bars represent SD). (C) Histogram showing the total number of B lymphocytes (B220+) and T lymphocytes (CD8+ or CD4+) in total bone marrow (collected from two hind legs) of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–8 mice per group; data are shown as mean; error bars represent SD). (D) Representative FACS plots showing B lymphocytes (B220+) in bone marrow of 12- to 15-month-old mice of the indicated genotypes. (E) Histogram showing the total number of thymocytes of 12- to 15-month-old mice of the indicated genotypes (n = 5 mice per group; data are shown as mean; error bars represent SD). (F) Representative FACS plots showing T cell development in thymus of 12- to 15-month-old mice of the indicated genotypes. Note that the percentage of CD8+, CD4+ (double positive) T cells (immature T cells) is reduced in G3, Exo1+/+ but rescued in G3, Exo1/ mice.

866 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

Figure 3. Exo1 Deletion Rescues Cell Proliferation and Reduces Apoptosis in Telomere-Dysfunctional Mice (A) Representative photographs of BrdU-stained crypts in the small intestine of 12- to 15-month-old mice of the indicated genotypes (magnification bars, 200 mm). (B) Histogram showing the percentage of BrdU- and PCNA-negative crypts in the small intestine of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD). (C) Representative photographs of apoptotic cells (green nuclei) in basal crypts (circled) of the small intestine of 12- to 15-month-old mice of the indicated genotypes (magnification bars, 200 mm). (D) Histogram showing the number of TUNEL-positive nuclei per basal crypts in the small intestine of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD).

meiotic defects resulting in testis atrophy and sterility in both male and female mTerc+/+ mice (Wei et al., 2003).

arrest appeared in intestinal crypts of 24-month-old G3 mTerc/, Exo1/ mice (Figures 3B and 3D).

Exo1 Deletion Reduces Cell Cycle Arrest and Apoptosis in Intestinal Crypts of Aging G3 mTerc/ Mice Impaired maintenance of high-turnover organs in telomere-dysfunctional mice has been linked to increased apoptosis and impaired cell proliferation (Lee et al., 1998). In agreement with previous studies (Choudhury et al., 2007), the rate of BrdU incorporation was reduced in basal intestinal crypts of aged G3 mTerc/ mice compared with that of mTerc+/+ mice (Figures 3A and 3B). Exo1 deletion partially rescued proliferation rates of intestinal basal crypts in G3 mTerc/, Exo1/ mice compared with G3 mTerc/, Exo1+/+ mice (n = 5 mice per group, p = 0.0041, Figures 3A and 3B). Elevated rates of apoptosis were present in basal crypts of 12- to 15-month-old G3 mTerc/ mice compared with age-matched mTerc+/+ mice (Figures 3C and 3D). Deletion of Exo1 reduced apoptosis in 12- to 15-month-old G3 mTerc/, Exo1/ mice compared with G3 mTerc/, Exo1+/+ mice (n = 5 mice per group, p = 0.001). An increase in apoptosis and cell cycle

Exo1 Deletion Does Not Rescue Telomere Dysfunction in Aging G3 mTerc/ Mice Studies in yeast have shown that defects in mismatch repair genes can promote telomerase-independent proliferation (Rizki and Lundblad, 2001). In mammalian cells, telomerase-independent immortalization of cells involves the activation of an alternative lengthening mechanism of telomeres (ALT), which is based on homologous recombination and characterized by an overall lengthening of telomeres and an increase in telomere length heterogeneity (Bryan et al., 1995; Chang et al., 2003). An analysis of telomere length in basal intestinal crypts (Figure 4A), splenic white pulp (data not shown), and bone marrow (Figure S1B) of 12- to 15-month-old mice revealed a reduction of the average telomere length and an increase in the frequency of cells with critically short telomeres in G3 mTerc/ mice compared with mTerc+/+ mice. This reduction in telomere length in G3 mTerc/ mice was not rescued by Exo1 deletion (Figure 4A and Figure S1B). We also did not see an increase of telomere length

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 867

Figure 4. Exo1 Deletion Does Not Rescue Telomere Function but Does Impair Formation of Chromosomal Fusions in mTerc/ Mice (A) The histograms show the distribution of telomere fluorescence intensities (TFI) over all analyzed nuclei of mTerc+/+, Exo1+/+ (n = 276 nuclei), mTerc+/+, Exo1/ (n = 160 nuclei), G3, Exo1+/+ (n = 242 nuclei) and G3, Exo1/ (n = 263 nuclei) mice. The red dotted lines indicate the mean value of each genotype. The black line marks the threshold of critically short telomeres (TFI < 200 arbitrary units). (B) Histogram of the rate of telomere-free ends per metaphase in bone marrow cells of 12- to 15-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD). (C) Representative photograph of a metaphase of a G3, Exo1/ mouse, with white arrow pointing to telomere-free ends (magnification bar, 4 mm). (D) Histogram of the rate of anaphase bridges per total number of anaphases in basal crypts of small intestine of 12- to 15-month-old mice of the indicated genotypes (n = 5 mice per group; data are shown as mean; error bars represent SD). (E) Representative photograph showing an anaphase bridge in basal crypts of small intestine (magnification bar, 20 mm). (F) Representative photograph of SKY analysis showing no translocations in a metaphase spread from bone marrow of 24-month-old G3 mTerc/, Exo1/ mice.

heterogeneity in G3 mTerc/, Exo1/. Together, there is no evidence of systemic activation of ALT that rescued survival and organ maintenance in G3 mTerc/, Exo1/ mice. As a direct marker of telomere dysfunction, telomere-free chromosome ends were present in bone marrow-derived metaphases of G3 mTerc/ mice but rarely in mTerc+/+ mice (Figures 4B and 4C). Exo1 deletion did not rescue the prevalence of telomere-free ends in G3 mTerc/ mice (Figures 4B and 4C). Instead there was a further increase in telomere-free ends in G3 mTerc/, Exo1/ mice, likely due to the abrogation of checkpoint responses (Figure 3). Exo1 Deletion Reduces Chromosomal Fusion and Does Not Increase Chromosomal Translocations in Telomerase Knockout Mice Telomere dysfunction induces chromosomal fusion by activation of DNA repair pathways (Maser et al., 2007b;

868 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

Smogorzewska et al., 2002). We analyzed the rate of anaphase bridges—a morphological correlate of chromosomal fusions (Kirk et al., 1997; Rudolph et al., 2001)—in the intestine of aging mice. There was a significant increase in anaphase bridges in G3 mTerc/, Exo1+/+ mice compared with those in age-matched wild-type mice, but Exo1 deletion rescued the formation of anaphase bridges in G3 mTerc/ mice (Figures 4D and 4E). Spectral karyotyping (SKY) analysis on metaphase spreads from bone marrow cells did not reveal recurrent chromosomal translocations in 12- to 15month-old G3 mTerc/, Exo1+/+ mice (data not shown). Notably, deletion of Exo1 did not result in any detectable increase in chromosomal translocations in bone marrow cells from 24-month-old G3 mTerc/, Exo1/ mice (Figure 4F, n = 4 mice). Together, these results indicated that Exo1 deletion reduced the formation of chromosomal fusions and did not increase chromosomal

instability in G3 mTerc/, Exo1/ mice with an extended lifespan. We did not detect a significant effect of Exo1 deletion on telomeric g-strand integrity in mouse embryonic fibroblasts (Figures S2A–S2D). Exo1 Deletion Prevents DNA Damage Accumulation and Reduces DNA Damage Signaling in Telomere-Dysfunctional Mice The formation of DNA damage foci represents the most upstream molecular event associated with DNA damage signal induction in response to telomere dysfunction (d’Adda di Fagagna et al., 2003; Satyanarayana et al., 2004). DNA damage foci containing gH2AX and 53BP1 were detected in basal intestinal crypts of 12- to 15month-old G3 mTerc/ mice (Figures 5A–5D), but not in age-matched mTerc+/+ mice (Figures S3A and S3B). Exo1 deletion reduced the accumulation of DNA damage foci in G3 mTerc/, Exo1/ mice (Figures 5A–5D). Costaining of gH2AX with TUNEL-positive, apoptotic cells revealed that DNA damage foci were present in nonapoptotic cells of aging G3 mTerc/ mice, indicating that the DNA damage foci staining did not reflect DNA breaks in apoptotic cells (Figure S3C). Only 14% of the total number of DNA damage foci (gH2AX or 53BP1) colocalized with telomeres (Figures S3D and S3E). These data indicated that Exo1 deletion impaired the accumulation of DNA damage, likely including extratelomeric DNA damage, in telomere-dysfunctional mice. Activation of DNA damage signaling in response to dysfunctional telomeres and DNA damage involves p53 and its downstream target, p21 (Chin et al., 1999; Brown et al., 1997; Choudhury et al., 2007). In 12- to 15-monthold G3 mTerc/, Exo1+/+ mice, nuclear expression of p53 was detected in 47%, and p21, in 9.5%, of basal intestinal crypts (Figures 5E–5H). p53 and p21 expression was not detected in mTerc+/+ mice (Figures 5F and 5H, Figures S4A and S4B). Both p53 and p21 upregulation was reduced in G3 mTerc/, Exo/ mice compared with that of G3 mTerc/, Exo+/+ mice (p < 0.001). Exo1 Deletion Impairs Checkpoint Induction and DNA Damage Signaling in Response to g-IR To analyze whether the effects of EXO1 were specific for telomeric DNA damage or reflected a more general role of EXO1 in response to DNA damage, we analyzed the role of EXO1 in response to different types of DNA damage. Exo1 deletion significantly rescued cell cycle arrest (Figures 6A and 6B, p = 0.01) and apoptosis (Figures 6A and 6C, p = 0.03) in basal intestinal crypts of g-irradiated mice (24 hr after 4Gy). Similarly, cultures of Exo1/ ear fibroblasts exhibited reduced checkpoint responses compared with Exo1+/+ ear fibroblasts in response to g-IR (Figures 6D and 6E and Figure S4C). Exo1 deletion also increased the resistance of cells treated with 6-thioguanine—a purine analog inducing DNA damage (Figure 6F); however, Exo1 gene status had no significant effect on the sensitivity of cultured cells in response to hydroxy

urea treatment (Figure S4D). Exo1 deletion did not affect the formation of gH2AX foci in g-irradiated fibroblasts (data not shown) or in the intestine of g-irradiated mice (Figure 6G). However, intestinal epithelial cells of irradiated Exo1/ mice, compared with those of irradiated Exo+/+ mice, showed a significant reduction in 53BP1 foci formation (Figure 6H) and reduced induction of p53 and p21 (Figures S4E and S4F). Exo1 deletion also reduced phosphorylation of CHK2 and p53 in g-irradiated fibroblasts (Figure 6I and Figures S4G and S4H). Of note, total protein levels of CHK2 and p53 did not differ between nonirradiated Exo1/ versus Exo1+/+ fibroblasts, indicating that Exo1 gene status by itself had no effect on total protein levels (Figure 6I and Figures S4G and S4H). Total protein levels of p53 were increased in g-irradiated Exo1+/+ fibroblasts (Figure 6I and Figures S4G and S4H), in agreement with the observation that phosphorylation of p53 stabilizes the protein in response to g-IR (Chen et al. 1994)—a response that was diminished in g-irradiated Exo1/ fibroblasts. A similar mechanism could explain the slight increase in total protein levels of CHK2 in response to g-IR in Exo1+/+ fibroblasts and the impaired increase of total CHK2 levels in Exo1/ fibroblasts. Exo1 Deletion Impairs Upstream DNA Damage Signal Induction Together, our results indicated that Exo1 deletion did not prevent gH2AX foci formation but did impair the induction of DNA damage signals at DNA breaks. Studies in yeast have shown that EXO1 generates ssDNA at dysfunctional telomeres, and the generation of ssDNA has been shown to induce DNA damage signals in yeast and mammalian cells (Garvik et al., 1995; Lee et al., 1998; Lydall and Weinert, 1995; Zou et al., 2003). To analyze whether EXO1 can generate ssDNA at DNA breaks in mammalian cells, we used the experimental system of laser-induced DNA breaks (Maser et al., 1997; Rogakou et al., 1999; Greenberg et al., 2006). Nondenaturing BrdU staining revealed a significant reduction in ssDNA formation in Exo1/ versus Exo1+/+ fibroblasts at laser-induced DNA breaks (Figures 7A and 7B). gH2AX staining at laser-induced DNA breaks was not affected by Exo1 gene status, but recruitment of Replication Protein A (RPA) to laserinduced breaks was reduced in Exo1/ versus Exo1+/+ fibroblasts (Figures 7C and 7D). Cell cycle profiles were similar in nonirradiated Exo1/ and Exo1+/+ fibroblasts, indicating that inherent differences in cell proliferation did not influence these results (Figures S5A and S5B). The generation of ssDNA and RPA binding at ssDNA leads to an activation of ATR (Namiki and Zou, 2006). Western blot analysis showed an impaired induction of phospho-ATR in g-irradiated Exo1/ versus Exo1+/+ fibroblasts (Figure 7E and Figure S5C). In addition, the formation of ATR foci was impaired in nuclei of g-irradiated Exo1/ versus Exo1+/+ ear fibroblasts (Figures 7F and 7G). Immunostaining did not reveal ATR foci in aging mTerc+/+ mice (Figure S5D). Notably, 12- to 15-month-old G3 mTerc/, Exo1+/+ mice showed nuclear foci of ATR in intestinal

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 869

Figure 5. Exo1 Deletion Prevents Accumulation of DNA Damage and Reduces DNA Damage Signaling in Telomere-Dysfunctional Mice (A) Representative photographs showing g-H2AX-positive cells (white arrows) in basal crypts of 12- to 15-month-old mice of the indicated genotypes (magnification bars, 20 mm). The inlet shows a nucleus containing g-H2AX foci at high-power magnification. (B) Histogram showing the percentage of g-H2AX-positive basal crypts in 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD). (C) Representative photographs showing 53BP1 foci (white arrows) in basal crypts of 12- to 15-month-old mice of the indicated genotypes (magnification bars, 20 mm). The inlet shows a nucleus containing 53BP1 foci at high-power magnification. (D) Histogram showing the number of 53BP1 foci per basal crypt in 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD). (E) Representative photographs showing p53-positive cells (white arrows) in basal crypts of 12- to 15-month-old mice of the indicated genotypes (magnification bars, 50 mm). (F) Histogram showing the percentage of p53-positive basal crypts in 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD). (G) Representative photographs showing p21-positive nuclei (red arrows) in basal crypts of 12- to 15-month-old mice of the indicated genotypes (magnification bars, 100 mm). (H) Histogram showing the percentage of p21-positive intestinal basal crypts of 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD).

870 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

basal crypts, but ATR foci formation was reduced in 12- to 15-month-old G3 mTerc/, Exo1/ mice (n = 4 mice per group, p = 0.001, Figures 7H and 7I). ATR foci appeared in nuclei of intestinal basal crypts of 24-month-old G3 mTerc/, Exo1/ mice (Figure 7I), indicating that EXO1-independent mechanisms can activate ATR in very old double knockout mice. DISCUSSION This study provides direct experimental evidence that EXO1 induces DNA damage responses in mammalian cells. Moreover, Exo1 deletion prevents the accumulation of DNA damage and prolongs lifespan of telomeredysfunctional mice. Exo1 Deletion Impairs the Induction of DNA Damage Signals The current study shows that Exo1 deletion impairs the induction of checkpoint responses and confers resistance of mammalian cells and organs to g-IR, telomere dysfunction, and 6-thioguanine treatment. The study points to the formation of ssDNA as one mechanism by which EXO1 amplifies the generation of DNA damage signals at DNA breaks. Previous studies on yeast have shown that the generation of ssDNA is required for DNA damage signal induction (Garvik et al., 1995; Lee et al., 1998; Lydall and Weinert, 1995; Zou et al., 2003). Similar mechanisms exist in mammalian cells (Zou et al., 2003). However, the molecular mechanisms leading to the formation of ssDNA in response to DNA damage are not completely understood. In yeast, EXO1 has been shown to induce ssDNA in response to telomere dysfunction (Maringele and Lydall, 2002). Our study provides, to our knowledge, the first evidence that the nuclease domain of EXO1 participates in the formation of ssDNA, RPA recruitment, and ATR activation at DNA double-strand breaks and in response to telomere dysfunction in mammalian cells. This mechanism likely contributes to aggravate phenotypes of telomere dysfunction in mTerc/ mice. Notably, Exo1 deletion did not rescue cell survival in response to Hydroxy urea (HU) treatment. A possible explanation is that HU induces ssDNA-dependent signaling independent of EXO1 processing, since HU induces single-stranded DNA damage by itself. Exo1 Deletion Impairs the Accumulation of DNA Damage in Telomere-Dysfunctional Mice Although EXO1 does not prevent gH2AX foci formation in response to g-IR and laser treatment, Exo1 deletion markedly reduced the accumulation of gH2AX foci in vivo in mTerc/ mice. These data indicate that dysfunctional telomeres induce EXO1-dependent processes that lead to an accumulation of further DNA damage. Cells having undergone replicative senescence due to telomere shortening display numerous gH2AX foci throughout the genome that do not colocalize with telomeric regions (Sedelnikova et al., 2004). It is possible that Exo1 deletion

reduces the secondary formation of DNA damage in response to telomere dysfunction by inhibiting chromosomal fusions and fusion-bridge-breakage cycles that create extratelomeric DNA damage in telomere-dysfunctional mice. In agreement with this hypothesis, Exo1 deletion reduces the number of anaphase bridges in telomere-dysfunctional mice (Figures 4D and 4E). Other, less obvious mechanisms might be involved. It has been shown that telomere dysfunction increases intracellular ROS levels in senescent cells (Passos et al., 2007). Although the exact mechanisms are not understood, it is possible that deletion of Exo1 also reduces the accumulation of ROS and thereby the accumulation of DNA damage in senescent cells. Therefore, it is conceivable that EXO1 has different, additive effects on DNA damage accumulation and DNA damage signal induction in telomeredysfunctional mice. Its role in DNA break processing could directly contribute to the induction of DNA damage signals. In addition, EXO1 could increase the generation of extratelomeric DNA damage in the context of chronic telomere dysfunction. An accumulation of extratelomeric DNA damage has been associated with organismal aging (Sedelnikova et al., 2004). Based on our study, it is intriguing to speculate that EXO1-dependent processes contribute to this phenomenon. Exo1 Deletion Rescues Organ Maintenance and Lifespan of Telomere-Dysfunctional Mice This study reveals that Exo1 deletion improves the maintenance of high-turnover organs and prolongs survival of telomere-dysfunctional mice. These findings indicate that EXO1-dependent mechanisms could represent therapeutic targets to improve regeneration and organ homeostasis in the context of telomere dysfunction. There is growing evidence that telomere dysfunction can impair organ function during human aging (Armanios et al., 2007; Cawthon et al., 2003; Tsakiri et al., 2007; Vulliamy et al., 2004) and chronic diseases (Ball et al., 1998; Wiemann et al., 2002). Since organ homeostasis is maintained by a balance of cell proliferation and apoptosis, it is likely that the inhibition of cell cycle arrest and apoptosis improved organ maintenance and lifespan of G3 mTerc/, Exo1/ mice compared with G3 mTerc/, Exo1+/+ mice. This interpretation of the data stands in agreement with our recent study that showed that deletion of p21 improves cell proliferation and organ maintenance in telomere-dysfunctional mice (Choudhury et al., 2007). The current study also shows that cell cycle arrest and apoptosis were not completely rescued in G3 mTerc/, Exo1/ mice; checkpoint activation and phenotypes of impaired organ homeostasis appeared in very old double knockout mice, but not in mTerc+/+ mice. These data suggest that mammalian cells possess EXO1-dependent and EXO1independent mechanisms for checkpoint induction in response to telomere dysfunction. Consistent with this notion, studies in yeast provide evidence that other exonucleases participate in the processing of dysfunctional telomeres (Tomita et al., 2003).

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 871

Figure 6. Exo1 Deletion Impairs DNA Damage Signal Induction in Response to g-IR (A) Representative photographs of BrdU-positive cells or TUNEL-positive cells in the intestinal basal crypts of 4-month-old mice of the indicated genotypes, 24 hr after g-irradiation (g-IR). Magnification bars, 200 mm. (B and C) Histogram showing the number of (B) BrdU-positive cells or (C) TUNEL-positive cells per crypt in the small intestine of 4-month-old g-irradiated mice of the indicated genotypes (n = 5 mice per group; data are shown as mean; error bars represent SD). (D and E) Cell cycle analysis in response to g-IR (24 hr after 15Gy). (D) Histogram showing the relative reduction of BrdU-positive cells in g-irradiated compared with nonirradiated ear fibroblasts of the indicated genotypes (n = 3 cell lines per group; data are shown as mean; error bars represent SD). (E) Histogram showing the relative increase in cells in G1 stage of the cell cycle in g-irradiated compared with nonirradiated ear fibroblasts of the indicated genotypes (n = 3 cell lines per group; data are shown as mean; error bars represent SD). (F) Survival curves of mouse embryonic stem cells of the indicated genotypes at 3–4 days after 6-thioguanine treatment at indicated doses. Survival curves for Exo1–/– (complete gene knockout) and Exo1exon6D/ (knockout of the nuclease domain, which was used throughout this study) were generated in two separate experiments. The survival curves for Msh2/ and WT are the averaged survival curves from two separate experiments. (G) Histogram showing the number gH2AX foci per nuclei in the small intestine of 4-month-old mice (n = 5 per group; data are shown as mean; error bars represent SD) of the indicated genotypes, 24 hr after 4Gy g-IR.

872 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

Exo1 Deletion Does Not Increase the Rate of Cancer Formation in Aging TelomereDysfunctional Mice The current study shows that Exo1 deletion did not increase cancer formation in telomere-dysfunctional mice. Studies in telomerase-deficient mice have shown that telomere dysfunction can increase tumor initiation by increasing chromosomal fusions and the rate of chromosomal instability (Artandi et al., 2000). Our study shows that Exo1 deletion impaired anaphase bridge formation in telomere-dysfunctional mice. These data are in agreement with the model that the generation of chromosomal fusions and consequent induction of fusion-bridge-breakage cycles represent a major mechanism inducing chromosomal instability and cancer initiation in the context of telomere dysfunction (Maser and DePinho, 2002; Zhu et al., 2003; O’Hagan et al., 2002; Maser et al., 2007a). The current study suggests that EXO1 contributes to the formation of chromosomal fusions and to the evolution of chromosomal instability as a consequence of fusionbridge-breakage cycles. In agreement with this interpretation, it has been shown that EXO1 mediates translocations at chromosome ends in response to telomere dysfunction in yeast (Hackett and Greider, 2003). In conclusion, our studies show that EXO1 is a critical component for inducing DNA damage signals, cell cycle arrest, and apoptosis in telomere-dysfunctional mice and in response to DNA double-strand breaks. Moreover, Exo1 deletion prevents the accumulation of DNA damage and can prolong organismal survival in the context of telomere dysfunction without accelerating chromosomal instability and cancer formation. EXPERIMENTAL PROCEDURES Mouse Crosses and Survival Exo1+/ mice were crossed with mTerc+/ mice to generate double heterozygous mice, which in turn were crossed through successive generations to produce G3 mTerc/, Exo1+/+ (n = 25) and G3, Exo1/ mice (n = 34). Intercrosses between mTerc+/+, Exo1+/ generated mTerc+/+, Exo1+/+ (n = 89) and mTerc+/+, Exo1/ mice (n = 27). Mice were on a C57BL/6J background. In Vivo BrdU Incorporation Assay In vivo proliferation was determined after 4 hr of BrdU labeling (i.p. injection of 30 mg/kg body weight). Immunostaining was performed on 5 mm thick paraffin sections using Cell Proliferation Kit (Amersham Biosciences) according to the manufacturer’s instructions. The number of BrdU-positive and BrdU-negative crypts was counted in 20 low-power fields (2003) and the percentage of BrdU-negative crypts was calculated. FACS Analysis For fluorescence-activated cell sorting analysis (FACS), thymus, spleen, and hind limb bones were dissected and single-cell sus-

pensions were stained with antibody cocktails (BD Pharmingen) for 15 min on ice. TUNEL Staining The rate of apoptosis was determined by TUNEL assay (In situ cell death detection kit, Roche, Mannheim, Germany) on 5 mm thick paraffin sections of small intestine. The number of apoptotic cells per crypt was counted in 20 low-power fields (2003) per mouse (n = 4–5 mice per group). Staining for gH2AX, ATR, 53BP1, p53, and PCNA Immunofluorescence was performed on 4 mm thick paraffin sections of small intestine. Sections were deparaffinized and rehydrated and permeabilized in 1 mM sodium citrate buffer. Primary antibodies were used either over night at 4 C or for 2 hr at room temperature: gH2AX (UpState, 1:750 dilution), ATR (Cell Signaling, 1:100), 53BP1 (Cell Signaling, 1:100), p53 (R&D, 1:50) and PCNA (Calbiochem, 1:50 dilution). The following secondary antibodies were used: gH2AX and PCNA: anti-mouse Cy3-Zymed (1:200 and 1:300 respectively); 53BP1: anti-rabbit Fitc-Zymed (1:150); p53: anti-goat Fitc-BD Pharmingen (1:100). p21 Staining For p21 staining 5 mm thick paraffin sections of small intestine were blocked with M.O.M. blocking reagent (Vector Labs) for 1 hr and then incubated with mouse anti-mouse p21 (Santa Cruz) overnight at 4 C. The subsequent steps to detect p21 signal was performed using M.O.M. Immunodetection Kit (Vector Laboratories, USA). ATR Staining on Mouse Ear Fibroblasts Mouse ear fibroblasts were grown on coverslips and cells were fixed in 4% formaldehyde, permeabilized with 0.2% triton, washed twice with PBS, and then treated with ATR antibody (1:500 dilution, Cell Signaling) for 2 hr. The secondary antibody (anti-rabbit Fitc-conjugated, Dianova) was used for 1 hr at room temperature. ATR-foci-positive cells were counted in 40 low-power fields (4003). Quantitative Fluorescence In Situ Hybridization, or qFISH qFISH was performed as described (Satyanarayana et al., 2003) on 5 mm thick paraffin sections of small intestine and methanol/aceticacid-fixed bone marrow cells, respectively. Telomere length was analyzed from the nuclei of the basal crypts and bone marrow using the TFL-Telo Software from Peter Lansdorp. SKY SKY was performed as described previously (Frank et al., 2004) and according to the manufacturer’s instructions (Applied Spectral Imaging, Ltd. [ASI]; Migdal HaEmek, Israel). Spectral images were acquired using an epifluorescence microscope equipped with an interferometer (SpectraCube ASI), a custom-designed optical filter, and the SkyView software (ASI). Western Blot Whole-cell extracts of mouse ear fibroblasts were obtained in RIPA lysis buffer. Protein was subjected to 10% SDS-PAGE and detected using antibody against phosphor-ATR (1:1000, Cell Signaling), Phospho-CHK2-T68 (1:1000, Abcam), phospho-p53-Ser15 (1:1000; Cell Signaling), b-Actin (1:1000, Santa Cruz), pan-specific anti-p53 (R&D), and pan-specific anti-CHK2 (Calbiochem).

(H) Histogram showing the number of 53BP1 foci per crypt in the small intestine of 4-month-old mice (n = 5 per group; data are shown as mean; error bars represent SD) of the indicated genotypes: 24 hr after 4Gy g-IR. (I) Representative western blots showing expression of phosphorylated CHK2 (T68), phosphorylated p53 (Ser15), total CHK2 (pan specific Ab), total p53 (pan specific AB), and b-actin as a loading control in g-irradiated and nonirradiated mouse ear fibroblasts of the indicated genotypes. Similar results were obtained in triplicate experiments using pools of three cell lines per experiment.

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 873

Figure 7. Exo1 Deletion Impairs Formation of ssDNA, RPA Recruitment, and ATR Activation in Response to Telomere Dysfunction and DNA Breaks (A) Representative photographs and (B) histogram showing the formation of ssDNA (stained by nondenaturing BrdU staining) at laser-induced DNA breaks (stained by a phospho-BRCA1-antibody) in fibroblasts of the indicated genotypes (n = 3 cell lines per group; data are shown as mean; error bars represent SD). (C) Representative photographs and (D) histogram showing recruitment of RPA to laser-induced DNA breaks in fibroblasts of the indicated genotypes. Note that gH2AX staining is independent of the Exo1 genotype, but recruitment of RPA is reduced in Exo1/ compared with Exo1+/+ fibroblasts. (E) Western blot showing phosphorylated ATR (Ser345) in protein lysates from 15Gy g-irradiated ear fibroblasts at 24 hr after IR. (F) Representative photograph of ATR foci formation in nuclei of g-irradiated (30 min after 2Gy) and nonirradiated ear fibroblasts of the indicated genotypes. (G) Histogram showing the percentage of ATR-foci-positive nuclei of g-irradiated (30 min after 2Gy) and nonirradiated ear fibroblasts of the indicated genotypes (n = 3 cell lines per group; data are shown as mean; error bars represent SD). (H) Representative photographs showing ATR foci in nuclei of cells in basal crypts of 12- to 15-month-old mice of the indicated genotypes (magnification bar, 20 mm). (I) Histogram showing the percentage of ATR-positive basal crypts in 12- to 15- and 24-month-old mice of the indicated genotypes (n = 4–5 mice per group; data are shown as mean; error bars represent SD).

874 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

Cytotoxicity Assays Cytotoxicity of 6-thioguanine and HU were determined by growth inhibition assays (Horton et al., 2000). Cells were exposed to 6-thioguanine and HU for 24 hr. After drug exposure, the plates were washed and cells were grown in drug-free DMEM growth media for 3–4 days. Cells were counted in triplicates using a lysis procedure for the 6-thioguanine experiments (Butler, 1984). For HU-treated cells, after 48 hr of treatment, cells were trypsinized and counted under a microscope and in a haemocytometer. Results are expressed in percent survival by comparing the number of surviving cells in drug-treated wells to the number of control cells in untreated wells.

Proliferation Assay of Cell Cultures Mouse ear fibroblasts were seeded in 10 cm dishes and grown overnight. Cells were then irradiated with 15Gy irradiation, and 24 hr later cells were trypsinized and fixed in 70% cold ethanol for analysis. BrdU (10 mM) was added 3 hr before harvesting the cells. BrdU staining was performed according to the manufacturer’s instructions using anti-Brdu (BD Pharmingen).

Laser-Generated DNA Double-Strand Breaks Laser-generated DNA double-strand breaks were generated using a P.A.L.M. MicroBeam laser (337 nm) microdissection system (Carl Zeiss MicroImaging, Inc.) as previously described (Greenberg et al., 2006; Lukas et al., 2003). Cells were grown on coverslips for 36–48 hr in media containing 10 mM BrdU (Sigma-Aldrich) prior to laser treatment. Laser stripes were performed on 70–100 cells per coverslip. Cells were returned to a cell culture incubator at 37 C for 15 min.

Immunofluorescence Following Laser Stripes Immunofluorescence following laser stripes was performed as previously described (Greenberg et al., 2006). In brief, paraformaldehydefixed cells were permeabilized in PBS/0.5% triton for 5 min on ice. Incubations with primary and secondary antibodies were carried out at 37 C for 20 min. Microscopy was performed on a Nikon Eclipse 80i integrated with ImageProTM 6.2 imaging software (Phase 3 Imaging Systems) which was used to generate deconvolved images.

Statistics Statistical analysis was done using Microsoft Excel and Graph Pad Prism software. Unpaired Student’s t test was used to generate the p values for the data sets. Survival curves were calculated using the method of Kaplan and Meier.

Supplemental Data The Supplemental Data for this article includes additional materials and methods, five figures, and one table, and can be found online at http://www.cell.com/cgi/content/full/130/5/863/DC1/.

ACKNOWLEDGMENTS K.L.R. was generously supported by the Deutsche Forschungsgemeinschaft (Heisenberg Professorship to K.L.R.: Ru 745/8-1, Ru745 4-1, KFO119, SFB 738, REBIRTH), the Deutsche Krebshilfe e.V. (102236-Ru 2), the Roggenbuck-Stiftung, the Wilhelm-Sander-Stiftung, and the Fritz-Thyssen Stiftung. A.B.C. and T.A.K. are supported by intramural research programs of the NIH and NIEHS. W.E. is funded by the NCI (R01CA093484). Received: October 5, 2006 Revised: May 17, 2007 Accepted: August 20, 2007 Published: September 6, 2007

REFERENCES Allsopp, R.C., Vaziri, H., Patterson, C., Goldstein, S., Younglai, E.V., Futcher, A.B., Greider, C.W., and Harley, C.B. (1992). Telomere length predicts replicative capacity of human fibroblasts. Proc. Natl. Acad. Sci. USA 89, 10114–10118. Armanios, M.Y., Chen, J.J., Cogan, J.D., Alder, J.K., Ingersoll, R.G., Markin, C., Lawson, W.E., Xie, M., Vulto, I., Phillips, J.A., 3rd., et al. (2007). Telomerase mutations in families with idiopathic pulmonary fibrosis. N. Engl. J. Med. 356, 1317–1326. Artandi, S.E., Chang, S., Lee, S.L., Alson, S., Gottlieb, G.J., Chin, L., and DePinho, R.A. (2000). Telomere dysfunction promotes nonreciprocal translocations and epithelial cancers in mice. Nature 406, 641–645. Ball, S.E., Gibson, F.M., Rizzo, S., Tooze, J.A., Marsh, J.C., and Gordon-Smith, E.C. (1998). Progressive telomere shortening in aplastic anemia. Blood 91, 3582–3592. Blackburn, E.H. (2001). Switching and signaling at the telomere. Cell 106, 661–673. Brown, J.P., Wei, W., and Sedivy, J.M. (1997). Bypass of senescence after disruption of p21CIP1/WAF1 gene in normal diploid human fibroblasts. Science 277, 831–834. Bryan, T.M., Englezou, A., Gupta, J., Bacchetti, S., and Reddel, R.R. (1995). Telomere elongation in immortal human cells without detectable telomerase activity. EMBO J. 14, 4240–4248. Butler, W.B. (1984). Preparing nuclei from cells in monolayer cultures suitable for counting and for following synchronized cells through the cell cycle. Anal. Biochem. 141, 70–73. Cawthon, R.M., Smith, K.R., O’Brien, E., Sivatchenko, A., and Kerber, R.A. (2003). Association between telomere length in blood and mortality in people aged 60 years or older. Lancet 361, 393–395. Chang, S., Khoo, C.M., Naylor, M.L., Maser, R.S., and DePinho, R.A. (2003). Telomere-based crisis: functional differences between telomerase activation and ALT in tumor progression. Genes Dev. 17, 88–100. Chen, C.Y., Oliner, J.D., Zhan, Q., Fornace, A.J., Jr., Vogelstein, B., and Kastan, M.B. (1994). Interactions between p53 and MDM2 in a mammalian cell cycle checkpoint pathway. Proc. Natl. Acad. Sci. USA 91, 2684–2688. Chin, L., Artandi, S.E., Shen, Q., Tam, A., Lee, S.L., Gottlieb, G.J., Greider, C.W., and DePinho, R.A. (1999). p53 deficiency rescues the adverse effects of telomere loss and cooperates with telomere dysfunction to accelerate carcinogenesis. Cell 97, 527–538. Choudhury, A.R., Ju, Z., Djojosubroto, M.W., Schienke, A., Lechel, A., Schaetzlein, S., Jiang, H., Stepczynska, A., Wang, C., Buer, J., et al. (2007). Cdkn1a deletion improves stem cell function and lifespan of mice with dysfunctional telomeres without accelerating cancer formation. Nat. Genet. 39, 99–105. d’Adda di Fagagna, F., Reaper, P.M., Clay-Farrace, L., Fiegler, H., Carr, P., Von Zglinicki, T., Saretzki, G., Carter, N.P., and Jackson, S.P. (2003). A DNA damage checkpoint response in telomere-initiated senescence. Nature 426, 194–198. de Lange, T. (2005). Shelterin: the protein complex that shapes and safeguards human telomeres. Genes Dev. 19, 2100–2110. Djojosubroto, M.W., Choi, Y.S., Lee, H.W., and Rudolph, K.L. (2003). Telomeres and telomerase in aging, regeneration and cancer. Mol. Cells 15, 164–175. Frank, O., Rudolph, C., Heberlein, C., von Neuhoff, N., Schrock, E., Schambach, A., Schlegelberger, B., Fehse, B., Ostertag, W., Stocking, C., and Baum, C. (2004). Tumor cells escape suicide gene therapy by genetic and epigenetic instability. Blood 104, 3543–3549. Garvik, B., Carson, M., and Hartwell, L. (1995). Single-stranded DNA arising at telomeres in cdc13 mutants may constitute a specific signal for the RAD9 checkpoint. Mol. Cell. Biol. 15, 6128–6138.

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 875

Greenberg, R.A., Chin, L., Femino, A., Lee, K.H., Gottlieb, G.J., Singer, R.H., Greider, C.W., and DePinho, R.A. (1999). Short dysfunctional telomeres impair tumorigenesis in the INK4a(delta2/3) cancer-prone mouse. Cell 97, 515–525. Greenberg, R.A., Sobhian, B., Pathania, S., Cantor, S.B., Nakatani, Y., and Livingston, D.M. (2006). Multifactorial contributions to an acute DNA damage response by BRCA1/BARD1-containing complexes. Genes Dev. 20, 34–46.

regional amplification and deletion in cancer genomes. Cancer Cell 2, 149–155. Passos, J.F., Saretzki, G., Ahmed, S., Nelson, G., Richter, T., Peters, H., Wappler, I., Birket, M.J., Harold, G., Schaeuble, K., et al. (2007). Mitochondrial dysfunction accounts for the stochastic heterogeneity in telomere-dependent senescence. PLoS Biol. 5, e110. 10.1371/ journal.pbio.0050110.

Hackett, J.A., and Greider, C.W. (2003). End resection initiates genomic instability in the absence of telomerase. Mol. Cell. Biol. 23, 8450–8461.

Qi, L., Strong, M.A., Karim, B.O., Armanios, M., Huso, D.L., and Greider, C.W. (2003). Short telomeres and ataxia-telangiectasia mutated deficiency cooperatively increase telomere dysfunction and suppress tumorigenesis. Cancer Res. 63, 8188–8196.

Hao, L.Y., Armanios, M., Strong, M.A., Karim, B., Feldser, D.M., Huso, D., and Greider, C.W. (2005). Short telomeres, even in the presence of telomerase, limit tissue renewal capacity. Cell 123, 1121–1131.

Rizki, A., and Lundblad, V. (2001). Defects in mismatch repair promote telomerase-independent proliferation. Nature 411, 713–716.

Hemann, M.T., Strong, M.A., Hao, L.Y., and Greider, C.W. (2001). The shortest telomere, not average telomere length, is critical for cell viability and chromosome stability. Cell 107, 67–77. Horton, J.K., Prasad, R., Hou, E., and Wilson, S.H. (2000). Protection against methylation-induced cytotoxicity by DNA polymerase betadependent long patch base excision repair. J. Biol. Chem. 275, 2211–2218. Kirk, K.E., Harmon, B.P., Reichardt, I.K., Sedat, J.W., and Blackburn, E.H. (1997). Block in anaphase chromosome separation caused by a telomerase template mutation. Science 275, 1478–1481. Lechel, A., Satyanarayana, A., Ju, Z., Plentz, R.R., Schaetzlein, S., Rudolph, C., Wilkens, L., Wiemann, S.U., Saretzki, G., Malek, N.P., et al. (2005). The cellular level of telomere dysfunction determines induction of senescence or apoptosis in vivo. EMBO Rep. 6, 275–281. Lee, H., Blasco, M., Gottlieb, G., Horner, J.W., 2nd, Greider, C., and DePinho, R. (1998). Essential role of mouse telomerase in highly proliferative organs. Nature 392, 569–574. Lukas, C., Falck, J., Bartkova, J., Bartek, J., and Lukas, J. (2003). Distinct spatiotemporal dynamics of mammalian checkpoint regulators induced by DNA damage. Nat. Cell Biol. 5, 255–260. Lydall, D., and Weinert, T. (1995). Yeast checkpoint genes in DNA damage processing: implications for repair and arrest. Science 270, 1488–1491. Maringele, L., and Lydall, D. (2002). EXO1-dependent single-stranded DNA at telomeres activates subsets of DNA damage and spindle checkpoint pathways in budding yeast yku70Delta mutants. Genes Dev. 16, 1919–1933.

Rogakou, E.P., Boon, C., Redon, C., and Bonner, W.M. (1999). Megabase chromatin domains involved in DNA double-strand breaks in vivo. J. Cell Biol. 146, 905–916. Rudolph, K.L., Chang, S., Lee, H.W., Blasco, M., Gottlieb, G.J., Greider, C., and DePinho, R.A. (1999). Longevity, stress response, and cancer in aging telomerase-deficient mice. Cell 96, 701–712. Rudolph, K.L., Millard, M., Bosenberg, M.W., and DePinho, R.A. (2001). Telomere dysfunction and evolution of intestinal carcinoma in mice and humans. Nat. Genet. 28, 155–159. Satyanarayana, A., Wiemann, S.U., Buer, J., Lauber, J., Dittmar, K.E., Wustefeld, T., Blasco, M.A., Manns, M.P., and Rudolph, K.L. (2003). Telomere shortening impairs organ regeneration by inhibiting cell cycle re-entry of a subpopulation of cells. EMBO J. 22, 4003–4013. Satyanarayana, A., Greenberg, R.A., Schaetzlein, S., Buer, J., Masutomi, K., Hahn, W.C., Zimmermann, S., Martens, U., Manns, M.P., and Rudolph, K.L. (2004). Mitogen stimulation cooperates with telomere shortening to activate DNA damage responses and senescence signaling. Mol. Cell. Biol. 24, 5459–5474. Sedelnikova, O.A., Horikawa, I., Zimonjic, D.B., Popescu, N.C., Bonner, W.M., and Barrett, J.C. (2004). Senescing human cells and ageing mice accumulate DNA lesions with unrepairable double-strand breaks. Nat. Cell Biol. 6, 168–170. Shay, J.W., and Wright, W.E. (2000). Hayflick, his limit, and cellular ageing. Nat. Rev. Mol. Cell Biol. 1, 72–76. Smogorzewska, A., Karlseder, J., Holtgreve-Grez, H., Jauch, A., and de Lange, T. (2002). DNA ligase IV-dependent NHEJ of deprotected mammalian telomeres in G1 and G2. Curr. Biol. 12, 1635–1644.

Maringele, L., and Lydall, D. (2004). EXO1 plays a role in generating type I and type II survivors in budding yeast. Genetics 166, 1641–1649.

Takai, H., Smogorzewska, A., and de Lange, T. (2003). DNA damage foci at dysfunctional telomeres. Curr. Biol. 13, 1549–1556.

Maser, R.S., and DePinho, R.A. (2002). Connecting chromosomes, crisis, and cancer. Science 297, 565–569.

Tomita, K., Matsuura, A., Caspari, T., Carr, A.M., Akamatsu, Y., Iwasaki, H., Mizuno, K., Ohta, K., Uritani, M., Ushimaru, T., et al. (2003). Competition between the Rad50 complex and the Ku heterodimer reveals a role for Exo1 in processing double-strand breaks but not telomeres. Mol. Cell. Biol. 23, 5186–5197.

Maser, R.S., Monsen, K.J., Nelms, B.E., and Petrini, J.H. (1997). hMre11 and hRad50 nuclear foci are induced during the normal cellular response to DNA double-strand breaks. Mol. Cell. Biol. 17, 6087–6096. Maser, R.S., Choudhury, B., Campbell, P.J., Feng, B., Wong, K.K., Protopopov, A., O’Neil, J., Gutierrez, A., Ivanova, E., Perna, I., et al. (2007a). Chromosomally unstable mouse tumours have genomic alterations similar to diverse human cancers. Nature 447, 966–971. Maser, R.S., Wong, K.K., Sahin, E., Xia, H., Naylor, M., Hedberg, H.M., Artandi, S.E., and DePinho, R.A. (2007b). DNA-dependent protein kinase catalytic subunit is not required for dysfunctional telomere fusion and checkpoint response in the telomerase-deficient mouse. Mol. Cell. Biol. 27, 2253–2265. Namiki, Y., and Zou, L. (2006). ATRIP associates with replication protein A-coated ssDNA through multiple interactions. Proc. Natl. Acad. Sci. USA 103, 580–585. O’Hagan, R.C., Chang, S., Maser, R.S., Mohan, R., Artandi, S.E., Chin, L., and DePinho, R.A. (2002). Telomere dysfunction provokes

876 Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc.

Tsakiri, K.D., Cronkhite, J.T., Kuan, P.J., Xing, C., Raghu, G., Weissler, J.C., Rosenblatt, R.L., Shay, J.W., and Garcia, C.K. (2007). Adult-onset pulmonary fibrosis caused by mutations in telomerase. Proc. Natl. Acad. Sci. USA 104, 7552–7557. Vaziri, H., and Benchimol, S. (1996). From telomere loss to p53 induction and activation of a DNA-damage pathway at senescence: the telomere loss/DNA damage model of cell aging. Exp. Gerontol. 31, 295–301. Vulliamy, T., Marrone, A., Szydlo, R., Walne, A., Mason, P.J., and Dokal, I. (2004). Disease anticipation is associated with progressive telomere shortening in families with dyskeratosis congenita due to mutations in TERC. Nat. Genet. 36, 447–449. Wei, K., Clark, A.B., Wong, E., Kane, M.F., Mazur, D.J., Parris, T., Kolas, N.K., Russell, R., Hou, H., Jr., Kneitz, B., et al. (2003). Inactivation of Exonuclease 1 in mice results in DNA mismatch repair defects,

increased cancer susceptibility, and male and female sterility. Genes Dev. 17, 603–614. Wiemann, S.U., Satyanarayana, A., Tsahuridu, M., Tillmann, H.L., Zender, L., Klempnauer, J., Flemming, P., Franco, S., Blasco, M.A., Manns, M.P., and Rudolph, K.L. (2002). Hepatocyte telomere shortening and senescence are general markers of human liver cirrhosis. FASEB J. 16, 935–942. Wong, K.K., Maser, R.S., Bachoo, R.M., Menon, J., Carrasco, D.R., Gu, Y., Alt, F.W., and DePinho, R.A. (2003). Telomere dysfunction and Atm deficiency compromises organ homeostasis and accelerates ageing. Nature 421, 643–648.

Wright, W.E., and Shay, J.W. (1992). The two-stage mechanism controlling cellular senescence and immortalization. Exp. Gerontol. 27, 383–389. Zhu, X.D., Niedernhofer, L., Kuster, B., Mann, M., Hoeijmakers, J.H., and de Lange, T. (2003). ERCC1/XPF removes the 30 overhang from uncapped telomeres and represses formation of telomeric DNAcontaining double minute chromosomes. Mol. Cell 12, 1489–1498. Zou, L., Liu, D., and Elledge, S.J. (2003). Replication protein Amediated recruitment and activation of Rad17 complexes. Proc. Natl. Acad. Sci. USA 100, 13827–13832.

Cell 130, 863–877, September 7, 2007 ª2007 Elsevier Inc. 877