Bacterial conjugation is a form of horizontal gene transfer that serves as an important source of genetic plasticity and diversity in both Gram positive and Gram negative bacteria. It has also been proved to be advantageous for dissemination of antibiotic resistance. Conjugation is a plasmid-mediated mechanism that allows single stranded DNA to be transferred across cell membranes from a donor to a recipient cell. In Escherichia coli, the conjugative F (fertility) plasmid encodes all necessary proteins for assembling the transfer apparatus thus allowing transfer and propagation of DNA into a recipient bacterial cell. Previous conflicting studies of bacterial conjugation in E. coli have shown DNA transfer to occur between donor and recipient pairs in intimate cell-cell contact or between distant donor and recipient pairs, connected only by the pilus, a filamentous protein extended from the donor cell. Using various fluorescent protein constructs to differentially mark donor and recipient cells and F plasmid DNA itself, we visualize conjugation in real time with single-cell resolution by fluorescent microscopy. Here we show that conjugation occurs only when donor and recipient cells are in intimate and close physical contact with each other. In addition, there appears to be no preference for cellular orientation for transfer. Furthermore, we propose a simple rate equation to model DNA transfer between bacteria and measure the rate of DNA transfer per unit time per cell contact. This quantitative model is a tool to elucidate conjugation at a single-cell scale and will provide a deeper understanding into the mechanism of bacterial conjugation.

All cells including bacteria exhibit an intricate spatial organization that is essential for cellular function. The mechanisms by which this ultra structural organization is established and maintained are not yet understood. The resolution limit of conventional wide-field fluorescence microscopy is set to 250nm by the diffraction limit, however the building blocks cellular ultra-structure, protein, are typically a few nanometers in size. Therefore, in order to probe the cell-machinery that gives rise to the organization of Escherichia coli chromosome, I use a super resolution fluorescence microscopy technique called PhotoActivation Localization Microscopy (PALM). I am able to resolve DNA structures with a resolution near ~20nm by labeling short sections of E. coli chromosome with a photoactivatable fluorescent protein. I then relate the measured size of chromosome sections to known physical parameters of DNA in order to asses the viability of various polymer models of DNA, which will lead to a better understanding of the physical factors that lead to structure in E. coli.

Sequencing the genome of an organism, or determining the order of the base pairs in its DNA, is one approach to understanding that organism’s chromosome structure.  Genome sequence assembly is accomplished with the help of computer programs, which often fail to include or map repetitious sequences in the final chromosome assembly.  One such unmapped sequence in the Lachancea waltii (L. waltii) yeast genome is the ribosomal DNA (rDNA), a sequence composed of hundreds of identical repeating segments that encodes the RNA component of ribosomes, the protein-producing machines in the cell.  My previous research has shown the rDNA is located in two large sequence gaps on chromosome 8, the only chromosome published incompletely as three separate segments of sequence.  My current research involves determining the order and orientation of these sequence scaffolds to verify and complete the assembly of L. waltii chromosome 8.  To investigate the arrangement of the three scaffolds, I am first performing indirect end labeling, where I am cleaving the chromosome with restriction enzymes to produce a specific fragmentation pattern.  Alignment of this pattern with that expected based on the proposed structure of chromosome 8 would validate the proposed chromosome structure and rDNA location.  In addition, I am performing a snap-back assay, a test that will indicate the orientation of the rDNA segment repeats – thereby verifying if the rDNA is present in one or two locations.  Finally, I am degrading the ends of chromosome 8 using an exonuclease – an end-specific enzyme – to determine the sequences, and thus the identity and orientation of the scaffolds, that are present at the chromosome ends.  By generating a complete, contiguous sequence of L. waltii chromosome 8, we hope to provide a more in-depth profile of this relatively uncharacterized yeast for future studies on chromosome evolution.

Centromeres in many species are found to reside near origins of replication that are activated early in S-phase, yet the biological significance of this conserved early replication of centromeres is unknown. Previous research has shown that the activation time of an origin is affected by its chromosomal location. A recent study has found that centromeres in the yeast Saccharomyces cerevisiae advance the activation time of nearby origins, and that this effect depends on the centromere’s ability to establish a functional kinetochore. These observations lead to the hypothesis that centromeres need to be replicated early to allow sufficient time for the assembly of functional kinetochores. If kinetochores fail to be established in a timely fashion, chromosomes may not be correctly bi-orient at mitosis. Daughter cells therefore may inherit the incorrect number of chromosomes creating conditions of aneuploidy such as those found in Down syndrome and many cancers. I will test the importance of early centromere replication by deleting origins immediately flanking a yeast centromere to delay that centromere’s replication. I will then conduct an assay to test for the rate of chromosome mis-segregation events in the mutant strain compared to the wild-type strain. I anticipate that the mutant strain may have a higher rate of loss for the modified chromosome, suggesting that early replication of centromeres is important for maintaining genome stability. This project will also provide insights into the regulation of the temporal program of replication origin activation and provide the first functional test for the consequences of disrupting the temporal program of chromosome replication.

Speciation is the process by which one species becomes two species that can no longer interbreed. A common hypothesis is that infertility between closely related species is due to the rapid evolution of DNA double-strand-break hotspots that are active in meiosis. DNA double-strand-breaks play an important role in meiosis by initiating recombination after chromosome replication and can be repaired as crossovers. However, hotspots where most of the breaks happen are preferentially lost to gene conversion during DNA repair. It is a mystery how hotspots are maintained over evolutionary time when they are constantly destroyed during meiosis. We hypothesize that hotspots rapidly appear and disappear over evolutionary time, which results in speciation. Schizosaccharomyces kambucha and Schizosaccharomyces pombe are two very closely related fission yeast species that are 99.5% identical at the DNA sequence level. Mapping out the hotspots of Schizosaccharomyces kambucha and comparing them to those of Schizosaccharomyces pombe will be a useful way to test our hypothesis. For this experiment, we have created Schizosaccharomyces kambucha mutants that can no longer repair the breaks so that we can pull out the broken DNA pieces using Chromatin Immunoprecipitation and locate where the DNA double-strand-break hotspots are in the genome. We expect to see the hotspots of Schizosaccharomyces kambucha differ from those of Schizosaccharomyces pombe, which will support the hypothesis that the difference in DNA double-strand-hotspots is indeed contributing to speciation. This research can be extended further to explain speciation in other organisms or unexplained infertility between human couples.

The process of cellular reproduction is dependent upon precise timing and coordination of regulatory events. Oscillations in cyclin-dependent kinase (Cdk) activities drive the cell cycle by regulating changes in the phosphorylation states of protein substrates. This dynamic and intricate signaling network of protein phosphorylation is a central component of the cell-cycle control system and enables the cell to modulate cellular functions and processes. Despite extraordinary advances in depth of coverage, phosphoproteomics experiments have struggled to expand in breadth, often comparing thousands of phosphorylation sites between only two moments in time. Such snapshots provide a biased view of the highly dynamic phosphoproteome and ignore important contextual information. To meet this challenge, we are developing robust and reproducible phosphoproteomic methodologies and utilizing mass spectrometry to study the changes in S. cerevisiae phosphoproteome over the course of the cell cycle. I first analyzed the reproducibility of the sample preparation techniques and data acquisition methodologies, which are key factors in the ability to distinguish true differences between different samples. Repeat analyses of 10 identical samples consisting of the entire yeast phosphoproteome were conducted, and preliminary results show that about 70% of the total identified phosphoproteins and 37% of the total identified phosphorylation sites were detected in all 10 repeat analyses with a variance in quantitation of less than 5%, indicating an accurate measurement of changes across the runs. I am now collecting global measurements of relative phosphorylation and protein changes in 5-minute intervals over the entirety of the cell cycle in a synchronously growing population to identify and quantify the proteins present at each time point of the cell cycle. This research will provide a wealth of information on the dynamics of proteome and phosphoproteome regulation throughout the cell cycle and unlock new insights into the biology of defective checkpoints in cancer.

Vitamin A, whose active form in our body is all-trans-retinoic acid (atRA), is essential for normal embryonic development and many other biological functions. CYP26A1 is an isoform of cytochrome P450 enzyme which is responsible for atRA metabolism and elimination. Since we cannot synthesize vitamin A on our own, humans must consume vitamin A as nutrients. Because both too much and too little intake of the vitamin are detrimental, a tightly regulated feedback system controls the level of vitamin A circulating in the body. CYP26A1, which is shown to be strongly inducible by atRA, takes a major role in modifying atRA into its inactive forms. Common aryl hydrocarbon environmental toxins are also known to induce the activity of CYP26A1 by binding to aryl hydrocarbon receptor (AHR). My current work aims to increase the understanding of the effects of environmental toxin on CYP26A1 in comparison to its other isoforms. I treated HepG2 cell line with varying concentration of 3-Methylcholanthrene (3MC) for 24 hours and quantified the amount of CYP26A1 mRNA. The results show that 3MC induces CYP26A1 transcription up to a certain toxic concentration. With better understanding of how common environmental toxins affect the health of an individual at a molecular level, it is possible to mediate the effects of the environmental toxins.