A‐type lamins are intermediate filament proteins that provide a scaffold for protein complexes regulating nuclear structure and function. Mutations in the LMNA gene are linked to a variety of degenerative disorders termed laminopathies, whereas changes in the expression of lamins are associated with tumourigenesis. The molecular pathways affected by alterations of A‐type lamins and how they contribute to disease are poorly understood. Here, we show that A‐type lamins have a key role in the maintenance of telomere structure, length and function, and in the stabilization of 53BP1, a component of the DNA damage response (DDR) pathway. Loss of A‐type lamins alters the nuclear distribution of telomeres and results in telomere shortening, defects in telomeric heterochromatin, and increased genomic instability. In addition, A‐type lamins are necessary for the processing of dysfunctional telomeres by non‐homologous end joining, putatively through stabilization of 53BP1. This study shows new functions for A‐type lamins in the maintenance of genomic integrity, and suggests that alterations of telomere biology and defects in DDR contribute to the pathogenesis of lamin‐related diseases.
Several studies have suggested that the 3D organization of the genome represents an additional level of regulation of genome function (Misteli, 2007; Dechat et al, 2008). The nuclear periphery, in particular, is viewed as a compartment actively participating in gene transcription and DNA repair (Therizols et al, 2006; Mekhail et al, 2008; Nagai et al, 2008). A‐type lamins, intermediate filament proteins that form part of the nuclear lamina and a nucleoplasmic network, are key determinants of nuclear architecture, providing a scaffold for the organization of nuclear functions (Taddei et al, 2004; Dechat et al, 2008). Accordingly, roles for A‐type lamins in DNA replication and repair, gene transcription and silencing, nuclear pore complex positioning, remodelling of chromatin, and nuclear envelope breakdown during mitosis have been suggested (Goldman et al, 2002; Gruenbaum et al, 2005). However, the causal relationship between alterations of A‐type lamins and regulation of genome functions as well as the molecular mechanisms involved remain ill defined. Elucidating the functions of these proteins is a topical subject, as mutations in the LMNA gene are associated with various degenerative disorders termed laminopathies (Broers et al, 2006; Capell and Collins, 2006). In addition, changes in the expression of A‐type lamins are associated with different types of human tumours (Agrelo et al, 2005; Prokocimer et al, 2006; Willis et al, 2008).
Most of the available data regarding A‐type lamins functions come from studies on human cells or mouse models carrying disease‐causing mutations in the LMNA gene, especially progerias (Mounkes and Stewart, 2004; Varela et al, 2005; Stewart et al, 2007). Common features include nuclear morphological abnormalities, disorganization of heterochromatin, and defects in the DNA damage response (DDR) pathway and in DNA repair (Liu et al, 2005; Capell and Collins, 2006). In particular, increased sensitivity to DNA damage has been molecularly linked to delayed recruitment of the DNA damage sensor 53BP1 to γ‐H2AX‐labelled DNA repair foci and defective formation of Rad 51 foci (Liu et al, 2005). Collectively, these data suggest that increased genomic instability contributes to the pathogenesis of disease‐causing mutations in the LMNA gene.
Increased genomic instability is also a known contributor to tumourigenesis. Although expression of mutant forms of A‐type lamins has not been associated with increased tumour susceptibility in humans or mice, an altered expression of A‐type lamins has been observed in different types of human tumours (Agrelo et al, 2005; Prokocimer et al, 2006, 2009; Willis et al, 2008). The LMNA knockout mouse has provided insights into the cellular consequences of the loss of A‐type lamins (Sullivan et al, 1999). Lmna−/− cells present alterations in the nuclear envelope, loss of heterochromatin from the nuclear periphery, and defects in the tethering and stabilization of members of two families of tumour suppressor proteins—the Retinoblastoma (Rb) and ING families—(Johnson et al, 2004; Han et al, 2008). Overall, these findings suggest common as well as different functional consequences of mutation versus silencing of the LMNA gene, with loss of A‐type lamins being associated with uncontrolled proliferation. The impact that the loss of A‐type lamins has on mechanisms responsible for maintaining genomic stability remains unknown.
Defects in DNA repair and the DDR pathway as well as alterations in telomere biology are among the leading causes of genomic instability in cancer and aging. Telomeres, heterochromatic structures sheltering the ends of linear chromosomes, are essential for the preservation of chromosome integrity and controlled cell proliferation (Blackburn, 2001; de Lange, 2002). A minimal length of telomeric DNA repeats and proper recruitment of telomere binding proteins are necessary to preserve telomere function (Liu et al, 2004; de Lange, 2005a). In addition, the acquisition of a heterochromatic structure at telomeres is crucial for the maintenance of telomere length homoeostasis (Blasco, 2007). The evidence linking A‐type lamins with aging, cancer, and at the cellular level, with heterochromatin regulation, has lead investigators to suggest that alterations in telomere biology could contribute to the phenotypes of laminopathies.
The best evidence about a role for A‐type lamins in telomere biology is that Hutchinson–Gilford progeria syndrome (HGPS) fibroblasts show faster telomere attrition than normal counterparts (Allsopp et al, 1992; Huang et al, 2008). HGPS fibroblasts also present defects in epigenetic marks characteristic of constitutive heterochromatin, such as decreased trimethylation of histone H3 at lysine 9 (H3K9me3) and increased trimethylation of histone H4 at lysine 20 (H4K20me3). However, the effect on specific domains, such as centromeres and telomeres is unknown (Scaffidi and Misteli, 2006; Shumaker et al, 2006). Further evidence of a crosstalk between A‐type lamins and telomeres is the finding that proliferative defects of human fibroblasts expressing lamin A mutants are rescued by telomerase (Kudlow et al, 2008), and that alterations of the nuclear lamina contributing to senescence are associated with aggregation of telomeres at the nuclear periphery (Raz et al, 2008).
To understand the role that A‐type lamins have in telomere biology, we determined the effect that loss of A‐type lamins has on the nuclear distribution of telomeres, and on the maintenance of telomere structure, length, and function. Using Lmna knockout mouse fibroblasts as a model (Sullivan et al, 1999), we provide evidence for an essential role of A‐type lamins in the 3D distribution of telomeres within the nucleus. Changes in the nuclear compartmentalization of telomeres upon loss of A‐type lamins were accompanied by telomere shortening, defects in telomeric chromatin structure, and increased genomic instability. In addition, we show that A‐type lamins are necessary for the processing of dysfunctional telomeres through the non‐homologous end joining (NHEJ) repair pathway. Interestingly, loss of A‐type lamins leads to a marked decrease in the cellular levels of 53BP1, a checkpoint protein that participates in the repair of double‐strand breaks, and that has been recently implicated in the processing of dysfunctional telomeres by NHEJ (Dimitrova et al, 2008). Overall, this study shows new functions for A‐type lamins in the maintenance of genomic stability through their participation in the maintenance of telomeres and 53BP1 stability. These findings suggest that alterations in telomere biology and defects in the DDR pathway are likely to contribute to the phenotypes of lamin‐related diseases, especially those characterized by the loss of A‐type lamins function.
Loss of A‐type lamins induces changes in the nuclear distribution of telomeres and telomere shortening
A 3D analysis of telomere positioning indicates that mammalian telomeres are distributed throughout the entire nuclear volume in G0/G1/S phases of the cell cycle, whereas they assemble into a telomeric disk at the centre of the nucleus during G2 (Chuang et al, 2004). The molecular mechanisms that ensure proper nuclear localization of mammalian telomeres and their relevance in telomere metabolism remain undefined. Interestingly, the 3D positioning of telomeres is altered in tumour cells (Chuang et al, 2004; Mai and Garini, 2006) and in senescent cells presenting defects in the nuclear lamina (Raz et al, 2008), suggesting a relationship between nuclear distribution of telomeres and alterations of telomere metabolism observed during senescence and immortality.
To determine whether A‐type lamins have a role in the nuclear compartmentalization of telomeres, we compared the nuclear distribution of telomeres between wild‐type (Lmna+/+) mouse embryonic fibroblasts (MEFs) and MEFs devoid of A‐type lamins (Lmna−/−). MEFs were analysed after spontaneously bypassing senescence and becoming immortal. We carried out 3D telomere fluorescence in situ hybridization followed by quantitative analysis to monitor telomere distribution (See Supplementary data). The telomere distances to the nuclear edge were determined using the TeloView program (see Vermolen et al, 2005 and Figure 1A). Figure 1B shows a clear difference in telomere distribution between the two genotypes, with a shift in the localization of telomeres towards the nuclear periphery upon loss of A‐type lamins. By calculating the cumulative distribution of telomere intensities (Figure 1C), we show that approximately 20% of telomere signals are found at the very edge of the nucleus (⩽0.4 μm) in both genotypes. However, although the remaining 80% of telomeres in Lmna−/− cells accumulate up to a distance of 1.75 μm from the edge, telomeres in Lmna+/+ MEFs expand throughout the nucleoplasm and up to the nuclear centre (2.75 μm). These results clearly show that A‐type lamins participate in the correct distribution of telomeres throughout the entire nuclear volume, with a significant change in distribution towards the nuclear periphery and away from the nuclear centre on loss of A‐type lamins. Fluorescence‐activated cell sorting analysis carried out on proliferating Lmna+/+ and Lmna−/− immortalized fibroblasts indicates that changes in telomere distribution are not due to differences in cell‐cycle profiles between genotypes (Supplementary Figure 1). In addition, chromatin immunoprecipitation (ChIP) assays performed using lamin A/C antibody show binding of lamins to telomeres (Supplementary Figure 2B), suggesting that tethering of telomeres to the lamins scaffold might regulate their nuclear distribution.
To determine whether changes in telomere distribution upon loss of A‐type lamins are accompanied by alterations of telomere metabolism, we compared telomere length between multiple sets of Lmna−/− and Lmna+/+ MEFs of early passage (pre‐senescent). On carrying out terminal restriction fragment (TRF) analysis (Garcia‐Cao et al, 2002), we observed a faster migration of telomeres in all five Lmna−/− lines compared with Lmna+/+ lines, indicating a moderate but highly consistent telomere shortening on loss of A‐type lamins (Figure 2A). These results were confirmed by quantitative fluorescence In situ hybridization (Q‐FISH) of metaphase nuclei using a telomeric probe (Garcia‐Cao et al, 2002). We found that mean telomere lengths of Lmna+/+ lines ranged between 37.6 and 40.8 kb, with an average telomere length of 39.0 kb. All five lines of Lmna−/− MEFs presented lower mean telomere length than any of the Lmna+/+ lines, ranging between 34.0 and 35.2 kb, with an average telomere length of 34.6 kb (Figure 2B). To determine whether these differences were statistically significant, we carried out a two‐sided t‐test considering the mean from each of the cell lines to be an independent sample of either Lmna−/− or Lmna+/+ genotype. We found that the mean telomere length is significantly different between these two genotypes (P=0.0003). In addition, TRF analysis of adult fibroblasts from Lmna−/− mice shows a more pronounced telomere shortening phenotype (Figure 2C). To test whether acute depletion of A‐type lamins would also lead to telomere attrition, we lentivirally transduced wild‐type MEFs with constructs carrying an shRNA specific for depletion of A‐type lamins (shLmna) or an shRNA specific for luciferase (shLucif) as control. Transduction using shLmna led to undetectable levels of lamins A/C (Figure 2D). Q‐FISH analysis carried out in these cells showed a marked decrease in telomere length after only five passages of the cells in culture (Figure 2E). All together, these data show that A‐type lamins have a key role in the maintenance of telomere length homoeostasis.
Telomere length homoeostasis is maintained primarily by telomerase (Blackburn et al, 2006). Furthermore, shelterin complex proteins, especially TRF1 and TRF2, regulate telomere length (Smogorzewska et al, 2000; Munoz et al, 2005). To further investigate the telomere shortening phenotype in Lmna−/− MEFs, we evaluated whether loss of A‐type lamins affects these regulatory mechanisms. We found no significant changes in telomerase activity between Lmna+/+ and Lmna−/− MEFs (Supplementary Figure 2A). In addition, ChIP analysis showed no detectable alterations in the telomere‐bound levels of TRF1 and TRF2 between genotypes (Supplementary Figure 2B). Our results indicate that the binding of these known regulators of telomere length homoeostasis is not affected by the loss of A‐type lamins. We cannot rule out, however, that the accessibility of telomerase to telomeres, or the binding of other factors implicated in telomere metabolism, especially factors implicated in telomere replication, are affected by the loss of A‐type lamins.
Loss of A‐type lamins results in alterations of telomere chromatin structure
The maintenance of a heterochromatic structure at telomeres is important for telomere length homoeostasis (Gonzalo et al, 2005, 2006). A common feature in fibroblasts from HGPS patients and from old individuals expressing progerin is the alteration of histone marks characteristic of constitutive heterochromatin (Scaffidi and Misteli, 2006; Shumaker et al, 2006), although the effect on telomeres is unknown. To investigate whether loss of A‐type lamins affects the assembly of telomeric heterochromatin, we carried out ChIP assays using antibodies recognizing well‐established heterochromatic marks, H3K9me3 and H4K20me3. Although we found no changes in H3K9me3 levels, we observed a significant decrease in telomeric H4K20me3 levels in Lmna−/− MEFs (Figure 3A). These results show that A‐type lamins participate in the maintenance of histone marks characteristic of telomeric heterochromatin. Interestingly, these defects were phenocopied by pericentric heterochromatin (Figure 3B), supporting the idea that alterations of A‐type lamins function affect the epigenetic status of constitutive heterochromatin (Shumaker et al, 2006). Nonetheless, the changes reported here are different from those described in HGPS cells, indicating different functional implications of mutation or silencing of the LMNA gene. The epigenetic defects of Lmna−/− MEFs were confirmed by western blot analysis, showing a marked reduction in global H4K20me3 levels with no changes in H3K9me3 (Figure 3C).
Recently, new components of telomeric heterochromatin have been identified. These are non‐coding telomeric RNAs (TERRAs or TelRNAs) that are transcribed by DNA‐dependent RNA polymerase II (Azzalin et al, 2007; Schoeftner and Blasco, 2008). Although the function of TERRAs on telomere metabolism remains unclear, different lines of evidence suggest that TERRAs might inhibit telomerase activity (Luke et al, 2008; Ng et al, 2009). The fact that alterations of A‐type lamins had been shown to impact RNA pol II‐directed transcription (Kumaran et al, 2002; Spann et al, 2002), prompted us to evaluate whether maintenance of TERRA levels is affected by their loss. We observed a marked decrease in the levels of TERRAs in Lmna−/− MEFs (Figure 3D), indicating that loss of A‐type lamins either inhibit RNA pol II‐directed transcription of TERRAs or alters their stability. All together, our data show that A‐type lamins have a role in the assembly of telomeric heterochromatin by stabilizing H4K20me3 and maintaining TERRA levels.
We had previously shown that stabilization of H4K20me3 at telomeres requires the action of histone methyltransferases, Suv4‐20h1 and Suv4‐20h2, as well as Rb family members (Gonzalo and Blasco, 2005; Benetti et al, 2007). In addition, Rb family members are transcriptional regulators, which could putatively modulate the transcription of TERRAs. Given that A‐type lamins contribute to the stabilization of Rb family members (Johnson et al, 2004), we hypothesized that the defects in telomeric chromatin structure on loss of A‐type lamins are due to Rb deficiency. We confirmed the decrease in the levels of Rb family members in our Lmna−/− MEF lines (Supplementary Figure 3), indicating that the lower levels of telomeric H4K20me3 are due, at least in part, to reduced Rb family function. We cannot rule out, however, that decreased Suv4‐20h1/h2 HMTase activity also contributes to this phenotype upon loss of A‐type lamins. Next, we tested whether the decrease in Rb family function could also be responsible for the reduction in TERRA levels on loss of A‐type lamins. We monitored TERRA levels in Rb family‐deficient MEFs (Rb−/−, Rb−/−p107−/− herein DKO, and Rb−/−p107−/−p130−/− herein TKO). Interestingly, we found increasing levels of TERRAs on loss of one, two, or three Rb family members (Supplementary Figure 4A). To control for differences in telomere length in Rb‐deficient MEFs (Supplementary Figure 4B), we normalized TERRA levels to telomeric DNA sequences (Supplementary Figure 4C). Remarkably, although the loss of A‐type lamins leads to decreased levels of TERRAs per telomeric DNA repeats, a similar increase in the density of TERRAs was observed in all Rb‐deficient MEFs (Figure 3E), indicating that reduced Rb function cannot account for the phenotype observed in Lmna−/− MEFs. Our findings thereby indicate that A‐type lamins play a role in the assembly of telomeric heterochromatin through Rb‐dependent and Rb‐independent mechanisms.
Intriguingly, Rb deficiency, decreased levels of H4K20me3, and decreased TERRA levels are consistent with a telomere elongation phenotype (Garcia‐Cao et al, 2002; Gonzalo et al, 2005; Benetti et al, 2007; Schoeftner and Blasco, 2008 and Supplementary Figure 4B). The fact that loss of A‐type lamins leads to telomere shortening despite Rb and H4K20me3 deficiency raises the possibility that A‐type lamins might have an active function in telomere elongation in these contexts. In summary, the alterations of telomere chromatin structure described here cannot account for the telomere shortening phenotype observed on loss of A‐type lamins. It is possible that a more global defect in chromatin structure and DNA metabolism because of alterations of nuclear architecture upon loss of A‐type lamins is responsible for this phenotype.
Impact of loss of A‐type lamins on telomere function and genomic stability
We evaluated whether the aforementioned changes in the nuclear distribution of telomeres, telomere length, and telomere chromatin structure translate into telomere dysfunction and genomic instability. We determined the loss of telomeric signals (signal‐free ends), the frequency of chromosome/chromatid breaks and end‐to‐end fusions, and the presence of aneuploidy in pre‐senescent Lmna−/− and Lmna+/+ MEFs (Figure 4 and Supplementary Figure 5). We found a threefold increase in the number of signal‐free ends (Figure 4A and Supplementary Figure 5A) and a twofold increase in breaks (Figure 4B and Supplementary Figure 5B) in Lmna−/− MEFs, indicating increased genomic instability. This finding was supported by an increased number of nuclei presenting basal DNA damage, as indicated by a twofold increase in cells presenting γ‐H2AX‐labelled foci upon loss of A‐type lamins (Figure 4C).
In normal cells, inappropriate recombination between telomeres of sister chromatids can serve to lengthen one telomere at the expense of another (Bailey et al, 2004; Laud et al, 2005). To test whether aberrant recombination involving telomeric repeats contributes to the loss of telomere signals in Lmna−/− MEFs, we carried out chromosome orientation fluorescence in situ hybridization (CO‐FISH) (Bailey et al, 2004). This technique allows the differential labelling of leading and lagging strands of telomeres. In the absence of recombination events between telomeric sequences, only one telomere at each chromosome end is labelled with either the leading or the lagging strand probe. If recombination occurs, the labelling is split between both sister telomeres giving rise to telomeres labelled with both the leading and lagging strand probes. CO‐FISH results indicate that loss of A‐type lamins does not lead to an increase in the frequency of recombination events involving telomeres (Supplementary Figure 6). Thus, alternative mechanisms are responsible for the increased loss of telomere signals upon loss of A‐type lamins.
In addition, karyotype analysis shows increased numbers of metaphases with abnormal chromosome dosage in Lmna−/− MEFs (Figure 4D). Earlier studies showed that reduced Rb family function leads to aneuploidy due, in part, to defects in centrosome duplication (Iovino et al, 2006). To test whether loss of A‐type lamins affects centrosome duplication control, we quantified centrosome numbers in Lmna+/+ and Lmna−/− MEFs by carrying out immunofluorescence using γ‐tubulin antibody. Our results indicate an increase in the number of cells presenting centrosome amplification (Figure 4E). Thus, loss of A‐type lamins leads to aneuploidy, at least in part, by deregulation of centrosome duplication cycle. Overall, these results show increased genomic instability on loss of A‐type lamins.
A‐type lamins participate in the processing of dysfunctional telomeres by NHEJ
Surprisingly, despite telomere shortening and increased frequency of signal‐free ends, we found no significant differences in the percentage of metaphases presenting chromosome end‐to‐end fusions in Lmna−/− MEFs (Supplementary Figure 5C). We hypothesized that loss of A‐type lamins might hinder the recognition and/or processing of dysfunctional telomeres through the NHEJ DNA repair pathway. To test this hypothesis, we carried out retroviral transduction of Lmna+/+ and Lmna−/− MEFs with a dominant‐negative mutant of the telomeric binding protein TRF2 (TRF2ΔBΔM) that induces telomere dysfunction and chromosome end‐to‐end fusions in the presence of an intact NHEJ repair pathway (Smogorzewska et al, 2002; Riha et al, 2006 and Figure 5A). Loss of telomere integrity on expression of TRF2ΔBΔM leads to the formation of DNA damage foci at telomeres, referred to as TIF (telomere dysfunction‐induced foci) (Takai et al, 2003; De Lange, 2005b). The ability of Lmna−/− MEFs to form TIF upon expression of TRF2ΔBΔM was evaluated by carrying out Immuno‐FISH using γ‐H2AX antibody and telomeric probe. The expression of TRF2ΔBΔM leads to the formation of γ‐H2AX‐labelled TIF in Lmna+/+ MEFs and Lmna−/− MEFs (Figure 5B and Supplementary Figure 7), suggesting that the initial step in the sensing of dysfunctional telomeres is intact on loss of A‐type lamins. In contrast, we found that the processing of dysfunctional telomeres by NHEJ is hindered by the loss of A‐type lamins. In Lmna+/+ MEFs, expression of TRF2ΔBΔM resulted in chromosome fusions in approximately 50% of metaphases, consistent with previous reports (Smogorzewska et al, 2002 and Figure 5C). Conversely, we found a fourfold reduction in the number of fusions in Lmna−/− MEFs expressing equivalent levels of TRF2ΔBΔM (Figure 5A and C). These results show that NHEJ of dysfunctional telomeres depends on the expression of A‐type lamins.
A‐type lamins stabilize 53BP1 protein levels
Similar to double‐strand breaks, dysfunctional telomeres activate the DDR pathway (Takai et al, 2003). Loss of A‐type lamins has been reported to affect the stability of nuclear factors, such as Rb family members and ING1 (Johnson et al, 2004; Nitta et al, 2006; Han et al, 2008). To gain insight into specific steps during DDR that could be affected by the loss of A‐type lamins hindering the processing of dysfunctional telomeres, we tested whether the cellular levels of key components of the DDR pathway could be altered in Lmna−/− MEFs. First, we monitored the levels H2AX, γH2AX, and 53BP1, which have a function in the sensing of dysfunctional telomeres (Takai et al, 2003). Although we found no differences in the levels of H2AX upon loss of A‐type lamins, we confirmed the increase in γH2AX levels in Lmna−/− MEFs, indicating increased basal DNA damage upon loss of A‐type lamins. In addition, we observed a marked decrease in the levels of 53BP1 in Lmna−/− MEFs when compared with Lmna+/+ cells (Figure 6A). The reduced levels of 53BP1 were confirmed by immunofluorescence. As shown in Figure 6B, the intensity of label with 53BP1 antibody is clearly reduced in Lmna−/− MEFs with respect to Lmna+/+under the same exposure conditions. We then tested whether the stability of other factors involved in DDR and in DNA repair are also affected by the loss of A‐type lamins. The cellular levels of MDC1, ATM, DNA‐PK, Mre11, Nbs1, Ku70, and ERCC1 were compared between Lmna+/+ and Lmna−/− MEFs (Figure 6C). We found no detectable differences in the levels of all these factors between the two genotypes. In addition, no changes in the global levels of proteins—TRF1, TRF2, and POT1—with a key structural function at telomeres were found (Figure 6D). These results indicate that the loss of A‐type lamins preferentially affects 53BP1. Interestingly, acute depletion of A‐type lamins by shLmna also significantly reduced the 53BP1 levels (Figure 6E). Most importantly, reintroduction of either lamin A, lamin C, or both lamins A/C into A‐type lamin‐depleted cells rescued the levels of 53BP1, demonstrating that lamins A and C are able to stabilize 53BP1 cellular levels.
To determine whether the decrease in the levels of this DNA damage sensor protein was the result of lower transcription, the levels of 53BP1 transcripts were monitored by quantitative PCR (Q‐PCR). As shown in Figure 6F, no differences in the levels of 53BP1 transcripts were detected between the two genotypes, indicating that the loss of A‐type lamins impacts the stability of the 53BP1 protein. Interestingly, incubation of Lmna+/+ and Lmna−/− MEFs with the proteasome inhibitor MG132 rescued the levels of 53BP1, implicating the proteasome pathway in the degradation of 53BP1 (Figure 6G). We hypothesize that similar to what has been reported for Rb family members (Johnson et al, 2004), A‐type lamins might stabilize 53BP1 protein levels by preventing their degradation by the proteasome. Loss of 53BP1 function has been recently associated with defective processing of dysfunctional telomeres by the NHEJ repair pathway. On the basis of our results we speculate that the decrease in 53BP1 levels could be responsible, at least in part, for the hindered ability of Lmna−/− MEFs to process dysfunctional telomeres.
Understanding the cellular functions of A‐type lamins is a highly topical subject because of their implication in a number of disease states, including laminopathies, aging and cancer. In particular, reduced expression of A‐type lamins is emerging as a factor contributing to tumourigenesis (Broers et al, 1993; Agrelo et al, 2005; Prokocimer et al, 2006). Our results indicate that A‐type lamins play a fundamental role in the maintenance of telomeres and genomic stability. We show that loss of A‐type lamins leads to a variety of alterations in telomere biology: (i) nuclear decompartmentalization of telomeres; (ii) impaired maintenance of telomere length homoeostasis; (iii) defects in telomere chromatin architecture; and (iv) a hindered ability to process dysfunctional telomeres by NHEJ. In addition, our results reveal a function for A‐type lamins in the DDR pathway, the stabilization of 53BP1 protein levels. Given that alterations of telomere biology and the DDR pathway are hallmarks of cancer and aging, we envision that the observed alterations in these processes, upon loss of A‐type lamins, are contributing to the pathogenesis of lamin‐related diseases, especially premature aging syndromes, such as HGPS, and tumoural processes characterized by the silencing of the LMNA gene.
Various lines of evidence indicate that the nucleus is compartmentalized and that changes in the spatial organization of chromatin affect nuclear functions (Goldman et al, 2002; Gruenbaum et al, 2005; Misteli, 2007). The importance of telomere compartmentalization for telomere function has been clearly shown in yeast (Akhtar and Gasser, 2007). To date, the mechanisms arranging the nuclear distribution of mammalian telomeres remain to be identified. In addition, how the nuclear localization of mammalian telomeres influences telomere biology is unknown. Lamins, which are absent in yeast, can bind directly to DNA and core histones, which attributes them a major role in tethering chromatin to specific sub‐nuclear compartments (Glass et al, 1993; Stierle et al, 2003). Our results show that A‐type lamins associate with telomeres and contribute towards their proper nuclear localization. The profound impact that loss of A‐type lamins has on different aspects of telomere biology suggests that the nuclear compartmentalization of telomeres could be fundamental for telomere metabolism. Our results do not indicate that changes in telomerase activity, the binding of TRF1 and TRF2, or aberrant recombination are the cause of the telomere shortening in Lmna−/− cells. It is possible that the accessibility of telomerase and/or activities participating in telomere metabolism is reduced on disruption of the scaffold for nuclear organization provided by A‐type lamins. Studies aimed to elucidate the impact of loss of A‐type lamins on telomere replication, and on the binding of other shelterin complex components or DNA repair factors to telomeres will be fundamental in understanding the mechanisms behind the alterations in telomere biology described here.
The change in localization of telomeres away from the nuclear centre and towards the periphery upon loss of A‐type lamins is intriguing. A‐type lamins are highly enriched at the nuclear periphery and are also found throughout the nucleoplasm (Schermelleh et al, 2008). We reasoned that loss of A‐type lamins could lead to the detachment of telomeres from the nuclear periphery. In contrast, we found that the localization of telomeres shifts towards the nuclear periphery in the absence of A‐type lamins, raising the possibility that the nuclear periphery might represent a default pathway for telomere localization. In this model, A‐type lamins would have an active role in the localization of telomeres throughout the nucleoplasm in mouse cells. A recent study showing that alterations of the nuclear lamina during senescence are associated with increased aggregation of telomeres at the nuclear periphery supports this model (Raz et al, 2008). It remains to be investigated whether tumour cells with silenced LMNA gene also present alterations in telomere compartmentalization and telomere structure, length, and function. These types of studies will provide insights into the mechanisms altered upon loss of A‐type lamins, which could contribute to tumourigenesis.
In addition to the effect on telomere biology, loss of A‐type lamins impacts on other molecular mechanisms, such as stabilization of Rb and ING tumour suppressors (Johnson et al, 2004; Nitta et al, 2006; Han et al, 2008) and the DNA damage sensor 53BP1 (this study). These mechanisms could contribute to the telomere phenotypes in Lmna−/− MEFs and to the genomic instability that drives cancer and aging. Our results show that indeed, reduced Rb family function is likely to be responsible for histone modifications defects in Lmna−/− cells. We show, however, that the telomere‐shortening phenotype and the decrease in TERRA density are Rb independent. Although the function of TERRAs in telomere biology is undefined, the observation that Rb family deficiency leads to increased levels of these new telomeric components is of great significance. Changes in TERRA density could contribute to the alterations in telomere metabolism previously described to occur upon loss of Rb family members. Future studies will need to address whether regulation of TERRAs and putatively, other non‐coding RNAs are part of the tumour suppressor function of Rb family.
Previous studies had shown that expression of mutant lamin A isoforms leads to alterations in the DDR and defective repair, which translate into increased sensitivity to DNA‐damaging agents (Liu et al, 2005; Scaffidi and Misteli, 2006; di Masi et al, 2008). To date, the effect that complete loss of A‐type lamins has on DNA repair mechanisms remains unknown. Our results show that the fusions resulting from dysfunctional telomeres upon expression of TRF2ΔBΔM require A‐type lamins, providing the first link between loss of A‐type lamins and defective NHEJ repair. The fact that mutations in the LMNA gene were previously associated with increased NHEJ (Liu et al, 2005) indicates yet another functional difference between mutation and loss of A‐type lamins. The increased genomic instability in Lmna−/− MEFs supports a role for A‐type lamins in DNA repair. In light of a recent study showing that loss of 53BP1 inhibits processing of dysfunctional telomeres by NHEJ (Dimitrova et al, 2008), we hypothesize that destabilization of this protein occurring upon loss of A‐type lamins is in part responsible for the observed phenotype. Similar to other nuclear factors, binding of 53BP1 to A‐type lamins either directly or through lamin‐associated proteins could contribute to its stability. The finding that loss of 53BP1 restricts the mobility of dysfunctional telomeres within the nucleus, lead to the proposal that 53BP1 has an active role in chromatin dynamics that facilitate the association and fusion of dysfunctional telomeres that might be far away within the nucleus (Dimitrova et al, 2008). We speculate that A‐type lamins could provide a platform for the association of 53BP1 with dysfunctional telomeres, as well as contribute to the mobility of dysfunctional telomeres to reach specific subcompartments where they come in contact with each other and with NHEJ repair factors.
Materials and methods
Lmna+/+ and Lmna−/− MEFs and adult fibroblasts were generated in the laboratory of Colin L Stewart as described by Sullivan et al (1999). Rb family‐deficient MEFs were generated in the laboratory of Julien Sage (Stanford University, CA). All lines were maintained in DMEM‐Glutamax (Invitrogen) supplemented with 10% bovine growth serum, antibiotics, and antimycotics. For cycloheximide and proteasome inhibitor treatments, 0.5 × 106 cells were cultured for 6 h in media containing 10 μg/ml cycloheximide, 30 μM MG‐132 (EMD Biochemicals) or EtOH as control.
ChIP analyses were carried out exactly as described by Garcia‐Cao et al (2004). Chromatin was immunoprecipitated using the following antibodies: anti‐H3K9me3 (#07‐442, Upstate); anti‐H4K20me3 (#07‐463, Upstate); anti‐TRF1 (T1948, Sigma); anti‐TRF2 (#05‐521, Upstate); or anti‐lamin A/C (SC‐6215, Santa Cruz). Immunoprecipitated DNA was slot‐blotted on a Hybond N+ membrane and hybridized to a telomeric probe (gift from T de Lange, Rockefeller University, NY, USA) or a major satellite probe. The quantification of the signal was done using the ImageQuant software (Molecular Dynamics). For input samples, serial dilutions of the unbound fraction of the no‐antibody control were processed. We calculated the amount of telomeric or pericentric DNA immunoprecipitated relative to the signal of the corresponding inputs. In all cases, we represented the ChIP values as a percentage of the total input telomeric DNA, therefore correcting for differences in the number of telomere repeats.
Immunoprecipitation, immunoblotting and immunofluorescence
Rb family members. Cells were lysed in RIPA buffer (0.15 M NaCl, 0.05 M Tris–HCl pH 7.2, 1% Triton‐X 100, 1% sodium deoxycholate, and 0.1% SDS), sonicated and 1 mg of total protein immunoprecipitated with antibodies bound to protein A‐agarose beads: Rb (IF18), p107 (C‐18), and p130 (C‐20) from Santa Cruz. Protein detection was carried out using antibodies against Rb (BD Pharmingen), p107 and p130 (Santa Cruz).
Lamin A/C, actin, γ‐tubulin, TRF2, and DDR factors. For immunoblotting, cells were lysed in RIPA buffer. Protein detection was carried out using Lamin A/C (SC‐6215, Santa Cruz), actin (Clone C4, MPB), β‐tubulin (Sigma), TRF1 (gift from Maria A Blasco), TRF2 (#05‐521, Upstate), POT1 (gift from Qin Yang), 53BP1 (Bethyl Laboratories, A300‐272A), MDC1 (gift from Junran Zhang), ATM (GTX7107, GeneTex), DNA‐PKcs (MS‐423‐P, NeoMarkers), Mre11 (Novus, MB100‐142), Nbs1 (Cell Signaling, 3002B), Ku70 (SC‐1486, Santa Cruz), H2AX (Upstate, 07‐164), γH2AX (Upstate, 07‐164), and ERCC1 (SC‐17809, Santa Cruz) antibodies. For immunofluorescence, cells were fixed using 4% paraformaldehyde and 0.1% Triton‐X 100 for 10 min at RT. Incubations using lamin A/C, γ‐tubulin (Sigma), and Alexa‐ or Cy3‐labelled secondary antibodies were done for 1 h. Washes were carried out in PBS and 20 mM glycine. Slides were counterstained using DAPI in Vectashield (Vector). Fluorescent images were taken using a Nikon 90i upright microscope or with a confocal microscope Zeiss L510.
Histones. Histones were purified by acid extraction as previously described by Shechter et al (2007). Histone modifications were detected by immunoblotting using anti‐H3K9me3 and anti‐H4K20me3 (Upstate).
Isolation of TERRAs
Total RNA and genomic DNA were isolated from the same samples using the TRI Reagent (Sigma) and following manufacturer's instructions. For RNA, 5 μg of isolated sample per well was separated in 1.2% agarose–formaldehyde gels, blotted on a Hybond N+ membrane and hybridized to telomeric and 28S probes. When indicated, negative controls were generated treating samples with RNase A (ABgene). For DNA, 0.1–0.5 μg of DNA was dot blotted on a nylon membrane and hybridized to telomeric or pericentric probes.
Telomerase activity was determined using the TeloTAGGG Telomerase PCR ELISA* Kit (Roche) following manufacturer's instructions.
Telomere length measurements
TRF analysis. We prepared cells in agarose plugs and carried out TRF analysis as described by Blasco et al (1997).
Q‐FISH on metaphases. We prepared metaphase stage chromosomal spreads for Q‐FISH and hybridized them as described by Herrera et al (1999). Fluorescent images were taken using a Nikon 90i upright microscope and intensity of telomere fluorescence analysed using the TFL‐Telo program (gift from P Lansdorp, Vancouver, Canada) (Zijlmans et al, 1997). Images and telomere fluorescence values were obtained from more than 50 metaphases in all cases.
Viral transduction with dominant‐negative TRF2, lamin A, lamin C, and shRNAs (shLmna and shLucif)
A dominant‐negative TRF2 mutant (TRF2ΔBΔM) cloned into the pLPC vector (gift from Titia de Lange, Rockefeller University) along with packaging and envelope plasmids pUMVC and pCMV‐VSV‐G (provided by Sheila Stewart, Washington University) were transfected in 293T packaging cells using Fugene. After 48 h post‐transfection, virus‐containing media was used to infect Lmna+/+ or Lmna−/− MEFs. Infection was repeated 24 h later. The same procedure was used to introduce lamins A/C into A‐type lamins depleted cells. Retroviral vectors for expression of lamins A/C were a gift from Brian Kennedy (University of Washington, Seattle, WA). GFP was used to monitor the efficiency of 293T transfection and MEF infection. After selection in puromycin (1 μg/ml) for 36–48 h, cells were prepared for FISH and western blot to test the expression of TRF2ΔBΔM. Lentiviral transduction of shLmna and shLucif cloned into pLKO was carried out in the same manner, using the packaging plasmid pHR’8.2ΔR and the envelope plasmid pCMV‐VSV‐G. Infected cells were subjected to immunoblotting after selection. Both shLmna and shLucif were a gift from Didier Hodzic (Washington University, St Louis, MO).
Cells growing in coverslips were first processed for immunofluorescence. Cells were fixed for 10 min in 3.7% paraformaldehyde/0.2% Tx‐100/PBS at RT, followed by three washes in PBS. After blocking in 10% BGS/PBS for 1 h, cells were incubated with anti‐γH2AX antibody 1 h at 37°C, washed three times and incubated with secondary antibodies. After washing extensively in PBS, cells were processed for FISH. Cells were fixed in 3.7% paraformaldehyde/PBS for 10 min at RT, washed in PBS, dehydrated in ethanol (70%, 90%, 100% ethanol, 5min each), and air dried. Hybridization solution containing the cy3‐labeled telomeric probe was added to coverslips, which were heated to 80°C for 10 min, and incubated for 3 h in the dark at RT. Coverslips were washed twice for 15 min in washing solution, and three times in PBS. Cells were counterstained with DAPI and coverslips mounted on slides.
CO‐FISH analysis was carried out as described by Bailey et al (2001). Briefly, cells were cultured in 20 μM BrdU for 22 h and 0.1 μg/ml colcemid was added for the last 4 h to enrich for mitotic cells. Metaphase stage chromosomal spreads were prepared as for Q‐FISH (Herrera et al, 1999). The slides were treated with RNase A (0.5 μg ml−1 for 10 min at 37°C, stained with Hoechst 33258 in 2 × SSC for 15 min at RT and exposed to 365 nm UV light for 30 min. Exonuclease III (3 U/μl) treatment for 20 min at RT was used to digest the BrdU‐labelled strands. The slides were then processed as for Q‐FISH but with two different probes added sequentially. Hybridizations were carried out first with the G‐rich probe (labelling the leading strand) followed by the C‐rich probe (labelling the lagging strand).
Quantitative real time PCR
Quantitative real‐time PCR was carried out on a MyiQ Detection system (BIO‐RAD, California, USA) using Taqman Universal PCR Master Mix (PE Applied Biosystems, California, USA). The cDNA was generated from 1 μg of total RNA using Geneamp RNA PCR kit (PE Applied Biosystems) following manufacturer's instructions. 53BP1 and GAPDH expression was determined using Taqman Gene expression assays (Mm00658689_m1 and Mm99999915_g1, respectively, PE Applied Biosystems). For the analysis, all reactions (in triplicate) were carried out by amplifying target gene and endogenous controls in the same plate. Relative quantitative evaluation of target gene was determined by comparing the threshold cycles.
A ‘two‐tailed’ Student's t‐test, ‘two‐samples of equal variance’ was used to calculate statistical significance of the observed differences in telomere length. Microsoft Excel v.2001 and Graphpad Instat v3.05 were used for the calculations. A paired t‐test to determine statistical significance was alternatively used when indicated. In all cases, differences were considered statistically significant when P<0.05.
Supplementary data are available at The EMBO Journal Online (http://www.embojournal.org).
Review Process File
We thank S Stewart, J Zhang and J Roti‐Roti of Washington University and J Davie of Manitoba Institute of Cell Biology for helpful discussions. We are grateful to T de Lange for providing TRF2ΔBΔM retroviral vector, D Hodzig for providing shLmna and shLucif vectors, and B Kennedy for the gift of lamins A/C expressing vectors. We thank A Searleman, L Revollo, K Chiappinelli, P Ramachandan, E Peterson, and C Franz for their contributions during their laboratory rotation. We are grateful to T Whitehead for his help with statistical analyses. CancerCare Manitoba Foundation supported the study carried out at the SM lab. Research in the laboratory of SG was supported by an institutional research grant from American Cancer Society, an Alzheimer's Disease Research Centre Pilot Grant, and Start‐Up funds provided by the Radiation Oncology Department at Washington University.
Author contributions: IGS and AR carried out most of the experiments and contributed towards writing this paper; SMP observed the decrease in 53BP1 levels. BV, DL, and SM carried out the 3D analysis of nuclear distribution of telomere intensity, and wrote the corresponding Supplementary data. DG carried out western blots of DDR factors. LM carried out CO‐FISH assays. EG and BPS carried out western blot analysis of ATM and DNA‐PK. JS provided all Rb family‐deficient MEFs. TS and CLS generated the Lmna−/− mouse model and provided the different lines of MEFs used in this study. SG supervised the research project and the preparation of the paper.
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