EEPD1 Rescues Stressed Replication Forks and Maintains Genome Stability by Promoting End Resection and Homologous Recombination Repair

Replication fork stalling and collapse is a major source of genome instability leading to neoplastic transformation or cell death. Such stressed replication forks can be conservatively repaired and restarted using homologous recombination (HR) or non-conservatively repaired using micro-homology mediated end joining (MMEJ). HR repair of stressed forks is initiated by 5' end resection near the fork junction, which permits 3' single strand invasion of a homologous template for fork restart. This 5' end resection also prevents classical non-homologous end-joining (cNHEJ), a competing pathway for DNA double-strand break (DSB) repair. Unopposed NHEJ can cause genome instability during replication stress by abnormally fusing free double strand ends that occur as unstable replication fork repair intermediates. We show here that the previously uncharacterized Exonuclease/Endonuclease/Phosphatase Domain-1 (EEPD1) protein is required for initiating repair and restart of stalled forks. EEPD1 is recruited to stalled forks, enhances 5' DNA end resection, and promotes restart of stalled forks. Interestingly, EEPD1 directs DSB repair away from cNHEJ, and also away from MMEJ, which requires limited end resection for initiation. EEPD1 is also required for proper ATR and CHK1 phosphorylation, and formation of gamma-H2AX, RAD51 and phospho-RPA32 foci. Consistent with a direct role in stalled replication fork cleavage, EEPD1 is a 5' overhang nuclease in an obligate complex with the end resection nuclease Exo1 and BLM. EEPD1 depletion causes nuclear and cytogenetic defects, which are made worse by replication stress. Depleting 53BP1, which slows cNHEJ, fully rescues the nuclear and cytogenetic abnormalities seen with EEPD1 depletion. These data demonstrate that genome stability during replication stress is maintained by EEPD1, which initiates HR and inhibits cNHEJ and MMEJ

Sensory Communication

Contains table of contents for Section 2, an introduction and reports on fourteen research projects.National Institutes of Health Grant RO1 DC00117National Institutes of Health Grant RO1 DC02032National Institutes of Health/National Institute on Deafness and Other Communication Disorders Grant R01 DC00126National Institutes of Health Grant R01 DC00270National Institutes of Health Contract N01 DC52107U.S. Navy - Office of Naval Research/Naval Air Warfare Center Contract N61339-95-K-0014U.S. Navy - Office of Naval Research/Naval Air Warfare Center Contract N61339-96-K-0003U.S. Navy - Office of Naval Research Grant N00014-96-1-0379U.S. Air Force - Office of Scientific Research Grant F49620-95-1-0176U.S. Air Force - Office of Scientific Research Grant F49620-96-1-0202U.S. Navy - Office of Naval Research Subcontract 40167U.S. Navy - Office of Naval Research/Naval Air Warfare Center Contract N61339-96-K-0002National Institutes of Health Grant R01-NS33778U.S. Navy - Office of Naval Research Grant N00014-92-J-184

A large genome-wide association study of age-related macular degeneration highlights contributions of rare and common variants.

This is the author accepted manuscript. The final version is available from Nature Publishing Group via http://dx.doi.org/10.1038/ng.3448Advanced age-related macular degeneration (AMD) is the leading cause of blindness in the elderly, with limited therapeutic options. Here we report on a study of >12 million variants, including 163,714 directly genotyped, mostly rare, protein-altering variants. Analyzing 16,144 patients and 17,832 controls, we identify 52 independently associated common and rare variants (P < 5 × 10(-8)) distributed across 34 loci. Although wet and dry AMD subtypes exhibit predominantly shared genetics, we identify the first genetic association signal specific to wet AMD, near MMP9 (difference P value = 4.1 × 10(-10)). Very rare coding variants (frequency <0.1%) in CFH, CFI and TIMP3 suggest causal roles for these genes, as does a splice variant in SLC16A8. Our results support the hypothesis that rare coding variants can pinpoint causal genes within known genetic loci and illustrate that applying the approach systematically to detect new loci requires extremely large sample sizes.We thank all participants of all the studies included for enabling this research by their participation in these studies. Computer resources for this project have been provided by the high-performance computing centers of the University of Michigan and the University of Regensburg. Group-specific acknowledgments can be found in the Supplementary Note. The Center for Inherited Diseases Research (CIDR) Program contract number is HHSN268201200008I. This and the main consortium work were predominantly funded by 1X01HG006934-01 to G.R.A. and R01 EY022310 to J.L.H

EEPD1 promotes replication fork restart after stress.

<p>(A) Replication recovery assayed as percentage of cells with ≥3 BrdU foci 2 h after release from 18 h treatment with 10 mM HU. Representative data (left) and quantitation (right) for cells transfected with control siRNA or si-EEPD1 targeted to 3’ UTR, with or without expression of siRNA-resistant FLAG-tagged EEPD1 (n = 11–23 determinations per condition, >100 cells scored/condition, means ±SEM). EEPD1 expression is shown by Western blot below for each condition. (B,C) Restart of stalled forks by DNA fiber analysis with HEK-293 cells transfected with empty vector (CMV), CMV-EEPD1 overexpression vector, control siRNA, or EEPD1 siRNA analyzed 15–30 min after release from HU replication stress. Representative images of fibers with IdU stained red and CldU stained green (B) and fiber quantitation (C) shown as percentage (means ±SD) of restarted forks (red + green fibers), stopped forks (red fibers), and new forks (green fibers) for >3 distinct determinations per condition (121–211 fibers/condition). (D) Fiber lengths and symmetry were measured in control and EEPD1 deficient cells to determine replication speed (left), and the percentage of bidirectional fibers (right), defined as red fibers with flanking green segments.</p

EEPD1 promotes end resection and downstream replication stress signaling.

<p>(A,B) End resection after IR measured by the fraction of cells with ss BrdU present in non-denatured DNA. Representative images (A) and quantitation (B) are shown as mean ±SD, n = 11–19 per condition, 139–180 cells/condition. (C) End resection adjacent to a single I-SceI DSB in HT1904 cells was measured in control cells and in cells depleted for EEPD1, CtIP and/or Exo1, alone or together (n = 3, means ± SD). For depletion of each protein, see <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1005675#pgen.1005675.s006" target="_blank">S6 Fig</a>. (D,E) Phospho-S4/S8 RPA32, gamma-H2Ax, and RAD51 nuclear foci formation in A549 cells after replication stress with and without EEPD1 depletion. DAPI nuclear counterstain is blue. (F,G) Phosphorylation of ATR (T1989), Chk1 (S345), and RPA32 (S4/S8) analyzed by Western blot in HU or mock-treated A549 cells with and without EEPD1 depletion. Representative blot (F) and quantification (n = 3–4 blots, means ±SD) (G) presented as relative protein levels normalized to β-actin loading controls. (H) Representative results (left) of alkaline single cell electrophoresis assays in untreated or HU-treated A549 cells with or with EEPD1 depletion, and quantitation (right) showing means ±SEM (n = 5).</p