Genomic translocations are well-established drivers of therapy-related myeloid neoplasms (t-MN) that affect survivors of primary malignancies. t-MN accounts for 10-20% of all myeloid malignancies and have very poor clinical outcomes. It is not possible to predict which patients treated for a primary cancer will develop t-MN, which constitutes a major clinical challenge. A method to assess the risk of translocations after patient exposures to DNA damaging chemotherapy and radiations would inform therapeutic decisions. Translocations depend on the movements of broken DNA ends on non-homologous chromosomes. We developed a method based on diffractive optical elements (DOE) to track photoactivated chromatin reporters (PACR) and map chromatin motions in the cell nucleus. Our preliminary data show that chromatin movements transiently decrease in response to DNA damage, which led to the hypothesis that chromatin `freezes' to facilitate the initial steps of the DNA damage response and reduce frequencies of genomic rearrangements. This effect may be mediated by reversible chromatin compaction and by chromatin interactions with structural nuclear elements. Inter-individual variability may determine genomic aberration frequencies and t-MN risk. The following specific aims will expand PACR assays and test this hypothesis:
In Aim 1, methodology will be developed for 3D measurements of chromatin mobility, in order to improve tracking accuracy and to account for inhomogeneities of the nuclear environment in all three dimensions. Rapid light sheet imaging will be used, and a novel multifocus 3D system based on a rotating point spread function DOE will be built. New photoactivatable DNA dyes will be developed to expand the PACR approach to difficult-to-transfect cells and tissues. The goals for Aim 2 are to identify the mechanisms controlling chromatin motions after DNA damage and to assess the functional consequences for DNA repair and translocations of chromatin `freezing'. We will feed PACR measurements with high time resolution (before/during/after DNA damage) into biophysical polymer models to predict physical changes of chromatin (intramolecular forces, persistence length, etc.), then test the predictions using multidimensional maps of chromatin motions, compaction, and nucleoskeletal organization, as well as functional cell assays. Two approaches to alter chromatin diffusion will assess causality between chromatin motions, DNA repair, and genomic translocations.
In Aim 3, we will use hematopoietic stem/progenitor cell samples from t-MN patients and healthy donors to test for clinically relevant associations between chromatin motions and genomic translocations. Characterizing the physical origins of genomic translocations may yield new methods to predict, and new targets to prevent, genomic rearrangements driving cancer initiation. Beyond the proposed research focused on chromatin motions during DNA repair, we anticipate broad applicability of our novel 3D imaging resources to study chromatin in multiple contexts.
Genomic translocations strongly increase the risk for leukemia, myelodysplastic syndromes, and lymphoma. The goal of this project is to further develop an optical method to quantify chromatin mobility after DNA damage and to characterize the relationship between chromatin motions and genomic translocations in cell models and patient samples. A method to assess the probability of genomic translocations will enable clinicians to predict and mitigate the risk of therapy-induced myeloid neoplasms, which is currently impossible.