Thursday, November 21, 2024
11/21/2024
Project Summary Single-molecule measurements of DNA and RNA, and their interactions and processing have revealed how these molecules function and how their activities are regulated. Our previous mechanistic studies of DNA unwinding enzymes and CRISPR enzymes have motivated us to develop novel biotechnological tools applicable in vitro and in living cells. We have also adopted next generation sequencing to develop high throughput biophysical assays for measuring DNA mechanics and chromatin remodeling activities in a massively parallel scale. We propose to pursue three broad areas of CRISPR-Cas9, chromatin remodeling, and DNA mismatch and its repair. A permeating theme is to explore the rich sequence space that is often poorly sampled in mechanistic studies. For effective genome editing using CRISPR-Cas9, Cas9 ribonucleoprotein complexes (RNPs) need to be assembled in high concentrations, requiring accurate co-transcriptional folding of the guide RNA and its stabilization by Cas9 binding. In addition, after target DNA cleavage, a Cas9-RNP needs to dissociate rapidly for the cell to detect the lesion and mount a DNA damage response. Understanding how these two critical steps are modulated by the target sequence will help researchers optimize target site selection and guide RNA design. Chromatin remodelers are ATPases that changes the position or composition of nucleosomes through nucleosome sliding or histone exchange, respectively. Many pioneering studies, including those using single- molecule methods, have revealed the fine details of the sliding reaction, but our knowledge of the histone exchange reaction is much more limited. We propose to develop single-molecule assays to fully define the kinetics and pathways of SWR1-catalyzed exchange of H2A-H2B dimer for H2A.Z-H2B dimer. We will also discover and characterize yeast native genomic sequences that allow nucleosome formation at a sharply defined location, which we hope will replace the artificial nucleosome positioning sequence, `601', that has been used in almost all mechanistic studies involving nucleosomes. DNA mismatch repair is carried out by conserved protein machinery but despite the decades of research, we have a very limited understanding of how different mismatches under different sequence contexts are repaired. For example, most mechanistic studies have been performed on a single type of mismatch (GT mismatch). We will develop a high throughput method to measure the in vivo mismatch repair efficiencies of thousands of different mismatches. If unrepaired until histone deposition on a nascent DNA, a mismatch will become part of a nucleosome. We will study the biophysical properties of a mismatch-containing nucleosome and its interplay with chromatin remodelers, allowing us to examine how an unrepaired mismatch can influence DNA accessibility.
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