The dimeric Mcm8–9 complex of Xenopus laevis likely has a conserved function for resistance to DNA damage

Most cancer treatments exploit the hypersensitivity of rapidly dividing tumor cells to DNA damage, largely reflecting problems with replicating damaged DNA templates. Many cancer chemotherapeutics directly damage DNA, and most types of DNA damage block replication forks. Other classes of chemotherapeutics include antimetabolites that reduce nucleotide pools and starve DNA polymerases or directly inhibit DNA polymerases, causing fork stalling. Blocked or stalled replication forks are initially stabilized by DNA damage response (DDR) proteins, including checkpoint and DNA repair proteins.1 Forks that fail to restart in a timely manner may regress to a “chicken foot” structure, which is subject to cleavage, causing fork collapse to double-strand breaks (DsBs).2 when replication forks encounter single-strand breaks (ssBs) and gaps (which can arise during repair of single-strand damage) this can result in direct fork collapse to DsBs. About half of cancer patients are treated with ionizing radiation, which directly induces DsBs, as well as base damage and ssBs that can be converted to DsBs during DNA replication. Thus, the common thread in all of these therapeutic strategies is DsB induction (Fig. 1). DsBs are highly cytotoxic, which explains their efficacy in cancer therapy and the intense effort to elucidate mechanisms of DsB induction and repair. several assays have been developed to measure DsB induction and repair. The induction of one or a few DsB at defined loci by the rare-cutting endonucleases i-scei and i-Ppoi, and their repair, can be measured with PCR assays using primers that flank the DsB.3 immunofluorescence microscopy is frequently used to detect phosphorylated histone Η2AX (γ-H2AX) foci, which appear adjacent to DsBs within 30 min of DsB induction, and their disappearance is taken as evidence of repair.4 γ-H2AX can also be detected by western blot, which provides an estimate of global DsB load in a population of cells. For more than 20 y, pulse field gel electrophoresis has been used to measure the fraction of broken DNA released from wells into the gel, providing a direct measure of DsBs in genomic DNA that is quantitative and reproducible. The comet assay is a related gel electrophoresis technique, in which DNA migrates out of individual cells embedded in agar on a microscope slide, producing DNA “tails” that extend from the body of the cell in a characteristic comet shape. Comet tail length (measured visually) and “tail moment” (product of tail length and the fraction of DNA in the tail determined by analysis of pixel intensities) are proportional to the number of DsBs; however, reproducible scoring of tail lengths or moments has proven difficult.5 each of the DsB assays above has its strengths and weaknesses, but none are particularly well-suited to high-throughput analysis. enter the engelward lab, which, in collaboration with engineers from the Bhatia lab, modified the comet assay to a 96-well format in which each of the 96 “macrowells” is subdivided into microfabricated “microwells,” ranging from 25–45 μm in diameter that each hold one to several cells.6 in a study by lead authors weingeist and Ge in the March 15, 2013 issue of Cell Cycle,7 engelward and colleagues at MiT and Harvard then demonstrated that this platform is very well-suited to high-throughput analysis of DsB induction and repair. The “CometChips” allow analysis of up to 96 different experimental conditions on a single gel, and because cells are arrayed, each comet can be scored using an automated image capture system, which greatly increases assay speed and reproducibility. How important is a reliable, high-throughput assay that directly measures DsB induction and repair? The DDR in general, and DsB repair in particular, are major determinants of cell survival and cell death and, thus, cancer treatment efficacy. The DDR is mediated by an incredibly complex network of proteins that includes, for example,