The chemistry of the genome is far more complex than the base pairing recognition of A-T and G-C. For example, small modifications to each base (e.g., DNA damage and epigenetic marks) profoundly impact a cell's ability to proliferate and produce secondary metabolites. On the other hand, direct binding of small molecules to DNA or RNA can be used to regulate biosynthetic pathways. Our research bridges the chemistry of the genome to natural product production and clinical diagnostics for new applications in green chemistry and cancer therapeutics.
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Strategies for mapping patterns of DNA damage in the genome are lagging behind gene sequencing/epigenetics because of the low abundance of DNA damage and because DNA damage impairs the enzymes used in sequencing methods. Recent developments include methods that use antibodies that bind DNA damage or high-fidelity enzymes that a amplify damage sites. Results from these pioneering methods support our hypothesis that DNA damage is not “random”; however, the methods cannot provide single-nucleotide resolution or lack specificity. To address this limitation, we recently developed “Click-Code-Seq”, a chemistry-based sequencing approach to precisely locate oxidative DNA damage. Click-Code-Seq relies on enzymes to remove specific damaged sites in the genome, followed by inserting a DNA “code” in its place. The DNA code can be sequenced and serves as an indicator for each position and type of damage. Our lab is developing new methods to sequence DNA damage and using these methods to characterize DNA damage patterns in the cancer genome to ultimately serve as a biomarker for chemotherapeutic resistance and sensitivity.
Fermentative production with Baker’s yeast, Saccharomyces cerevisiae, has garnered significant interest for the sustainable production of biofuels and natural products. For example, there has been a recent explosion of successful yeast-based preparation of important pharmaceuticals including fungal (e.g., penicillin) and plant (e.g., morphine) natural products as well as chemical building blocks/substrates for the production of biopolymers, plastics, and biofuels. Despite these successes, several roadblocks currently prevent yeast biosynthesis from becoming a wide-spread industrial alternative for chemical production. For example, importing multi-step biosynthetic pathways into yeast often requires balancing enzyme expression levels, minimizing pathway bottlenecks, and preventing self-toxic effects. Furthermore, the diversity of possible end-products is limited to natural biosynthetic pathways and their corresponding enzymatic reactions. Our group is harnessing nucleic acid technology (e.g., aptamers, ribozymes, nanoparticles) to resolve these biosynthetic limitations.
The last few years have seen rapid development of technologies that permit a detailed analysis of the genome and transcriptome of a single cell; however, corresponding techniques for defining single cell levels of metabolites are not available. Without these tools, it is difficult to link a specific perturbed cellular response to the quantitative internal dose, including the absorption, distribution, metabolism and excretion (ADME) of drugs, intracellular metabolites, toxic exposure, etc. As such, the McKeague lab is dedicated to the development of non-invasive, in vivo biosensors that report the concentration of metabolites inside specific cells. We are leveraging aptamer to develop these biosensors and are exploring their application in a wide range of model systems including yeast, bacteria, mammalian cells, and zebrafish.
