SummaryATR controls chromosome integrity and chromatin dynamics. We have previously shown that yeast Mec1/ATR promotes chromatin detachment from the nuclear envelope to counteract aberrant topological transitions during DNA replication. Here, we provide evidence that ATR activity at the nuclear envelope responds to mechanical stress. Human ATR associates with the nuclear envelope during S phase and prophase, and both osmotic stress and mechanical stretching relocalize ATR to nuclear membranes throughout the cell cycle. The ATR-mediated mechanical response occurs within the range of physiological forces, is reversible, and is independent of DNA damage signaling. ATR-defective cells exhibit aberrant chromatin condensation and nuclear envelope breakdown. We propose that mechanical forces derived from chromosome dynamics and torsional stress on nuclear membranes activate ATR to modulate nuclear envelope plasticity and chromatin association to the nuclear envelope, thus enabling cells to cope with the mechanical strain imposed by these molecular processes.

Cellular senescence, an irreversible proliferation arrest evoked by stresses such as oncogene activation, telomere dysfunction, or diverse genotoxic insults, has been implicated in tumor suppression and aging. Primary human fibroblasts undergoing oncogene-induced or replicative senescence are known to form senescence-associated heterochromatin foci (SAHF), nuclear DNA domains stained densely by DAPI and enriched for histone modifications including lysine9-trimethylated histone H3. While cellular senescence occurs also in premalignant human lesions, it is unclear how universal is SAHF formation among various cell types, under diverse stresses, and whether SAHF occur in vivo. Here, we report that human primary fibroblasts (BJ and MRC-5) and primary keratinocytes undergoing replicative senescence, or premature senescence induced by oncogenic H-Ras, diverse chemotherapeutics and bacterial cytolethal distending toxin, show differential capacity to form SAHF. Whereas all tested cell types formed SAHF in response to activated H-Ras, only MRC-5, but not BJ fibroblasts or keratinocytes, formed SAHF under senescence induced by etoposide, doxorubicin, hydroxyurea, bacterial intoxication or telomere attrition. In addition, DAPI-defined SAHF were detected on paraffin sections of Ras-transformed cultured fibroblasts, but not human lesions at various stages of tumorigenesis. Overall, our results indicate that unlike the widely present DNA damage response marker γH2AX, SAHF is not a common feature of cellular senescence. Whereas SAHF formation is shared by diverse cultured cell types under oncogenic stress, SAHF are cell-type-restricted under genotoxin-induced and replicative senescence. Furthermore, while the DNA/DAPI-defined SAHF formation in cultured cells parallels enhanced expression of p16(ink4a) , such 'prototypic' SAHF are not observed in tissues, including premalignant lesions, irrespective of enhanced p16(ink4a) and other features of cellular senescence.

Both Myc and Ras oncogenes impact cellular metabolism, deregulate redox homeostasis and trigger DNA replication stress (RS) that compromises genomic integrity. However, how are such oncogene‐induced effects evoked and temporally related, to what extent are these kinetic parameters shared by Myc and Ras, and how are these cellular changes linked with oncogene‐induced cellular senescence in different cell context(s) remain poorly understood. Here, we addressed the above‐mentioned open questions by multifaceted comparative analyses of human cellular models with inducible expression of c‐Myc and H‐RasV12 (Ras), two commonly deregulated oncoproteins operating in a functionally connected signaling network. Our study of DNA replication parameters using the DNA fiber approach and time‐course assessment of perturbations in glycolytic flux, oxygen consumption and production of reactive oxygen species (ROS) revealed the following results. First, overabundance of nuclear Myc triggered RS promptly, already after one day of Myc induction, causing slow replication fork progression and fork asymmetry, even before any metabolic changes occurred. In contrast, Ras overexpression initially induced a burst of cell proliferation and increased the speed of replication fork progression. However, after several days of induction Ras caused bioenergetic metabolic changes that correlated with slower DNA replication fork progression and the ensuing cell cycle arrest, gradually leading to senescence. Second, the observed oncogene‐induced RS and metabolic alterations were cell‐type/context dependent, as shown by comparative analyses of normal human BJ fibroblasts versus U2‐OS sarcoma cells. Third, the energy metabolic reprogramming triggered by Ras was more robust compared to impact of Myc. Fourth, the detected oncogene‐induced oxidative stress was due to ROS (superoxide) of non‐mitochondrial origin and mitochondrial OXPHOS was reduced (Crabtree effect). Overall, our study provides novel insights into oncogene‐evoked metabolic reprogramming, replication and oxidative stress, with implications for mechanisms of tumorigenesis and potential targeting of oncogene addiction.