Genes encoding the RNA portion of the ribosome (rDNA) are present in essentially all eukaryotic genomes as tandem repeated arrays. In humans, rDNA copy number is highly variable and an undervalued potential source of genetic disease. Changes in rDNA copy number can occur through DNA breakage and repair as well as through errors in DNA replication. High transcriptional activity at the rDNA locus poses challenges for replication; all tested eukaryotes have evolved replication fork barriers (RFBs), ensuring that replication machinery does not collide with transcribing RNA polymerases. In yeast, the RFB is a specific sequence to which the protein Fob1 binds, blocking replication forks that converge with transcription. Mutants lacking Fob1 have greatly reduced variation in rDNA copy number. There are currently two models to explain how Fob1 binding to the RFB produces rDNA copy number instability. One model suggests that binding of Fob1 actively recruits DNA break and repair machinery which induces recombination between rDNA repeats. Another model proposes that the stalled replication fork at the RFB is inherently fragile, increasing the likelihood of breakage. To distinguish between these two models, I am generating yeast strains where Fob1 binds to the RFB but does not arrest forks. Using CRISPR/Cas9 gene editing technology, I am reversing the direction of the RFB in each of the 150 rDNA repeats in yeast to prevent replication fork stalling. By confirming the absence of replication fork stalling and determining whether rDNA instability has also been reduced, I can distinguish whether Fob1 binding to the RFB in the absence of fork blocking contributes to rDNA copy number changes. Clarifying involvement of the RFB in rDNA copy number changes will additionally provide insights into the connections between transcriptional activity, replication fork stalling and genome instability.
 

Synthetic CRISPR–Cas transcription factors enable the construction of complex gene expression programs, and chemically–inducible systems allow for precise, rheostatic-like control over the transcriptional dynamics. We have constructed a bio-orthogonal, chemically–inducible synthetic CRISPR regulatory system to activate and repress gene expression in yeast. By fusing chemically inducible transcriptional regulators to specific RNA binding proteins, we have expanded the tunability of this system of constructs. The RNA binding proteins are fused to half of one chemically-inducible system while the other half is fused to an effector. Upon addition of drug, the two halves come together to form the complete chemically-inducible system and either activate or repress the target gene. We use reporter gene assays to probe the dose-dependence, time-dependence, and reversibility of the systems. The use of multiple, orthogonal chemically-inducible systems and unique guide RNAs allows for more sophisticated, multi-gene programs that still maintain precise control of gene expression dynamics independently at different sites.

New techniques in genome mapping have revealed a high degree of 3D spatial organization in the nucleus. Long-range loops connect enhancers to their gene targets to regulate expression. In order to assess the mechanisms and functions behind 3D spatial organization of the nucleus we need a system that allows us to engineer DNA loops. We use programmable CRISPR-Cas DNA binding domains to target specific sites in the genome. The CRISPR-Cas complex is tethered to a targeting domain (TD) that can dimerize with another TD at a distant DNA locus. To promote interactions between TDs that are bound to DNA, we have designed an allosteric switch that assembles the TD only when the CRISPR-Cas complex has engaged its DNA target. To validate that our switch proteins can act as a DNA sensor we have developed a simple reporter assay; upon successful switch protein activation on DNA a transcription factor is recruited to drive expression of a fluorescent protein. Our results indicate that the protein switches activate when they are recruited to DNA, effectively acting as a sensor for DNA binding. Future steps include optimizing our reporter assay design and modifying the design for DNA looping.

Hybrid vigor or heterosis describes superiority of a hybrid compared to its parents; however, the genetic mechanisms underlying this phenotype remain largely unresolved. One potential mechanism is loss of heterozygosity (LOH), a process where there is loss of one copy of a gene or surrounding chromosomal region. In our previous work, we evolved hybrids between two species of yeast, Saccharomyces cerevisiae and Saccharomyces uvarum - which differ in their temperature preference - by growing them in a chemostat at different temperatures in phosphate limited media for several hundred generations. We repeatedly observed LOH events in these hybrids in response to changes in temperature. Each LOH event incorporated the gene region encoding the Pho84 membrane bound inorganic phosphate transporter protein on chromosome 13, which is important when phosphate is limited in the environment. These repeated LOH events all affect fitness based on environmental temperature; however the events span various lengths with some as short as a few kilobases and others as large as 200 kilobases. Because they are different in length we also know that they include different numbers of genes. To investigate whether these varying LOH lengths may include other genes that affect hybrid fitness, we have used CRISPR/Cas9 to create double strand breaks at specific sites along the S. cerevisiae chromosome 13, resulting in DNA repair using the S. uvarum chromosome as a template and formation of LOH events of different lengths. Our aim is to create a pool of hybrids with varying LOH tracts and let them compete in a phosphate limited environment to assess the relationship between the different LOH regions and fitness. This will allow us to narrow down genes that may be responsible for temperature sensitivity or that contribute to higher hybrid fitness in organisms that are heterozygous or homozygous.

Different phenotypes of normal cells may influence genetic and epigenetic profiles and tumorigenicities of their transformed cells. The current study presents a comprehensive analysis of mutations in the whole mitochondrial genome for different immortalized (pre-neoplastic) cell types, which were derived from normal breast epithelial cells with two different phenotypes (stem vs non-stem). Duplex Sequencing, which shows the lowest error rates among currently available DNA sequencing methods, was applied to investigate rare and subclonal (low-frequency) mutations as well as homoplasmic (high-frequency) mutations. We found that the vast majority of mutations in the cells occur at low-frequency and are not detectable by conventional DNA sequencing methods. The most prevalent mutation types are C>T/G>A and A>G/T>C transitions. The overall frequencies of rare and subclonal mutations are significantly lower in stem cell-derived immortalized human breast epithelial cells (SV1) than in non-stem cell-derived immortalized human breast epithelial cells (SV22). The decreased mitochondrial mutagenesis of SV1 cells occurred mainly in the noncoding RNA region (rRNA and tRNA) of the mitochondrial genome. In addition, the predicted pathogenicity for rare and subclonal mutations in the tRNA region was significantly lower in SV1 cells than in SV22 cells. Our findings suggest that different phenotypes of normal cells lead to distinctive mitochondrial mutation profiles of immortalized cells. The identified mutation spectra and mutations specific to stem (versus non-stem)-derived immortalized cells have implications in characterizing heterogeneity of tumors and understanding the role of mitochondrial mutations in immortalization and transformation of human cells.

During embryonic development, a dormancy-like state known as diapause arises during the transition from pre to post implantation. This state of suspended development is a reproductive strategy which favors newborn survival in mammals during nutritional deprivation or stress. Studies from the Ruohola-Baker lab found potential candidate regulators of diapause by establishing an in-vitro diapause model using pluripotent mouse embryonic stem cells (mESC). One of the genes is Activating Transcription Factor 5 (ATF5) which encodes a protein capable of survival-mediated functions such as maintaining mitochondrial activity during stress, modulating cell differentiation, preventing apoptosis and regulating cancer pathway. ATF5 has been known to transcriptionally target mTOR, a mechanistic target of rapamycin. Energy stress in the form of starvation and pharmacological inhibition of mTOR has shown to induce diapause-like state in mESCs in vitro. Our hypothesis is that upregulation of ATF5 under energy stress will reestablish diapause-like state in naïve mouse embryonic stem cells in vitro. We will test our hypothesis by loss-of-function and overexpression experiments. We test if ATF5 gene knockout using CRISPR-Cas9 prevents the mutant lines from entering diapause-like state from energy stress. Using western blots, we will quantify phospho-mTOR levels and its downstream targets in the ATF5 KO lines and compare them with the wildtype lines. For the overexpression of ATF5, we will make rescue lines for the ATF5 KO cells. We predict that overexpressed ATF5 in rescue lines will enter diapause-like state, and have reduced mTOR and its downstream target signals compared to KO lines. Our discoveries of ATF5 function in diapause can be useful in understanding how early-staged cancer stem cells enter a diapause-like state or quiescent state which enables them to escape chemotherapy detection. We can potentially contribute to the development of therapies to target ATF5 mechanism so that these undetected cancer stem cells can be detected.

Recent studies have shown just how important microbiomes are for individual health, population health, and environmental health. Unfortunately, these studies are often limited by the costs of meta-Genomics. Furthermore, meta-Genomic data itself is limiting by only providing information on population characteristics, but not the functional contributions of members within the population. Single cell transcriptomic sequencing aims to lessen the latter issue by providing information on the gene expression of each individual cell within a sample. Even so, current single cell sequencing technologies are costly and require specialized equipment. SPLiT-seq is a single cell transcriptomic technology developed by the Seelig lab at the University of Washington that uses split-pool ligation to create uniquely barcoded cDNA for each cell using every-day laboratory bench tools and techniques and costs only one cent per cell. Currently, SPLiT-seq is well-optimized for mammalian cells. However, using this method on bacteria requires its own set of optimized procedures given the morphological and biochemical differences between eukaryotes and prokaryotes. The aims of this project are to deal with these biological differences to increase the information obtained from messenger RNA and decrease the amount received from ribosomal RNA, as well as reduce the amount of cells that get the same cDNA barcodes. By optimizing this single-cell transcriptomic technique for bacteria, future studies involving microbial communities will be able to obtain more robust information on the individuals within those populations.

In order for multipotent stem cells to properly differentiate into specialized cells, specific genes must be expressed at a specific time and amount during development. Many of the factors that regulate expression have been identified; however, it remains unclear how they work together to control the timing and amplitude of gene expression. Non-coding DNA elements, known as enhancers, can increase the the likelihood of transcription of a gene by integrating signals in the cell to provide regulatory logic for gene regulation. To understand how enhancers tune gene expression timing and amplitude during development, our lab has generated a transgenic mouse in which each of the two copies of the T-cell identity gene, Bcl11b, have been tagged with distinguishable fluorescent reporters, providing a sensitive readout for gene activity at the single locus level. Bcl11b turns on during T-cell development, and its activation executes a developmental switch from a hematopoietic stem cell to a T-cell committed progenitor. There is a non-coding region far downstream of Bcl11b which harbors a cluster of putative enhancers. To interrogate the function of individual candidate enhancers, we use CRISPR/Cas9 targeting to generate specific genomic deletions in T cell progenitors. From our preliminary experiments, we have shown that cutting off the entire enhancer region completely inhibits the expression of Bcl11b entirely compared to when we cut out only an individual enhancer peak which only partially inhibits it. This is promising because it shows that we have found an enhancer that controls the probability of activation while not being necessary for the activation of Bcl11b. This work will reveal the cis-regulatory logic that underlies the control of a master lineage-specifying gene. This better understanding will help us identify new strategies to control the expression of master regulatory genes for cellular reprogramming.