Browsing by Author "MacAlpine, David M"

Metabolism is known to be driven by intrinsic genetic programs as well as contextual factors within the environment. Individual genetic and environmental determinants of metabolic state have been extensively characterized, both within normal physiological processes as well as in the context of disease states such as cancer. However, it is becoming increasingly appreciated that the inevitable interaction between these differential sources of metabolic regulation can dramatically influence cellular phenotypes, a phenomenon commonly referred to as gene-environment interaction. These interactions can create substantial heterogeneity between individuals, particularly in the context of tumor metabolism which can ultimately impede the development and efficacy of many clinical therapies. Characterization of these relationships can therefore improve the predictive applicability of targeted therapeutic approaches, as well as contribute to the identification of novel treatment strategies that can circumvent the biological limitations imposed by gene-environment interactions. Using metabolomic, genetic, and pharmacological approaches, in this dissertation I examine the metabolic consequences of environmental alterations in defined genetic settings. I provide in-depth characterization of the relative predictability of cellular responsiveness to nutrient availability in the context of genetic deletion of the metabolic enzyme MTAP, results of which demonstrate potential implications in previously-identified metabolic vulnerabilities in MTAP-deleted cancers. I additionally examine how perturbation of energetic demand, via either pharmacological inhibition of the Na+/K+ ATPase or with the physiological stimulus of exercise, impacts metabolic processes in diverse biological contexts. This work collectively illustrates the exceptional heterogeneity in metabolic adaptation to environmental alterations, and provides support for the future development of lifestyle modifications and repurposing of common pharmacological agents as therapeutic modalities in cancer treatment.

The accurate and timely replication of eukaryotic DNA during S-phase is of critical importance for the cell and for the inheritance of genetic information. Missteps in the replication program can activate cell cycle checkpoints or, worse, trigger the genomic instability and aneuploidy associated with diseases such as cancer. Eukaryotic DNA replication initiates asynchronously from hundreds to tens of thousands of replication origins spread across the genome. The origins are acted upon independently, but patterns emerge in the form of large-scale replication timing domains. Each of these origins must be localized, and the activation time determined by a system of signals that, though they have yet to be fully understood, are not dependent on the primary DNA sequence. This regulation of DNA replication has been shown to be extremely plastic, changing to fit the needs of cells in development or effected by replication stress.

We have investigated the role of chromatin in specifying the eukaryotic DNA replication program. Chromatin elements, including histone variants, histone modifications and nucleosome positioning, are an attractive candidate for DNA replication control, as they are not specified fully by sequence, and they can be modified to fit the unique needs of a cell without altering the DNA template. The origin recognition complex (ORC) specifies replication origin location by binding the DNA of origins. The S. cerevisiae ORC recognizes the ARS (autonomously replicating sequence) consensus sequence (ACS), but only a subset of potential genomic sites are bound, suggesting other chromosomal features influence ORC binding. Using high-throughput sequencing to map ORC binding and nucleosome positioning, we show that yeast origins are characterized by an asymmetric pattern of positioned nucleosomes flanking the ACS. The origin sequences are sufficient to maintain a nucleosome-free origin; however, ORC is required for the precise positioning of nucleosomes flanking the origin. These findings identify local nucleosomes as an important determinant for origin selection and function. Next, we describe the D. melanogaster replication program in the context of the chromatin and transcription landscape for multiple cell lines using data generated by the modENCODE consortium. We find that while the cell lines exhibit similar replication programs, there are numerous cell line-specific differences that correlate with changes in the chromatin architecture. We identify chromatin features that are associated with replication timing, early origin usage, and ORC binding. Primary sequence, activating chromatin marks, and DNA-binding proteins (including chromatin remodelers) contribute in an additive manner to specify ORC-binding sites. We also generate accurate and predictive models from the chromatin data to describe origin usage and strength between cell lines. Multiple activating chromatin modifications contribute to the function and relative strength of replication origins, suggesting that the chromatin environment does not regulate origins of replication as a simple binary switch, but rather acts as a tunable rheostat to regulate replication initiation events.

Taken together our data and analyses imply that the chromatin contains sufficient information to direct the DNA replication program.

DNA replication is an intricate process within eukaryotic cells that must be precisely executed to preserve genetic information. This process begins at multiple start sites, or origins of replication, along each chromosome which are selected, licensed, and activated through cell-cycle regulated steps. Powerful reconstitution studies have identified the proteins involved in these processes, but they do not fully recapitulate the nuclear environment. Within the nucleus, the genome is organized in a chromatin structure consisting of DNA and all associated factors. At origins of replication, local chromatin contributes to origin identity and activation, but the precise chromatin dynamics that occur at these sites during helicase activation and initial DNA unwinding have not been fully explored. Additionally, how these steps are regulated to ensure genomic stability remain unstudied within the context of chromatin.

To address these questions, I have developed a conditional system that removes polymerase α function to capture helicase activation at replication origins in the budding yeast. Under restrictive conditions, these cells (cdc17-ts-FRB) do not initiate replication. When allowed to recover, replication appears to initiate outside origins, necessitating a delay in G2/M phase to repair unreplicated gaps at origins. To investigate origin chromatin and helicase movement prior to replication, I used MNase chromatin profiling alongside ChIP-seq for various replication factors. Chromatin in a 1 kb region around early, efficient replication origins is disrupted under restrictive conditions. The active helicase unwinds DNA out to 1 kb from these origins and is likely the source of the chromatin disruption. I next used the cdc17-ts-FRB conditional system to investigate the regulation of helicase progression in the absence of replication. I first tested whether the intra-S-phase checkpoint had a role in stalling the helicase 1 kb from the origin. Though removing checkpoint activation distributed helicase movement and chromatin disruption to late, inefficient origins, it did not alter the distance the helicase progressed from the origin. Instead, the helicase stalls as it leaves the AT-rich origin region and encounters sequences with higher GC content. These results provide in vivo support for the recently proposed “dead man’s switch” model for decreased helicase processivity when uncoupled from replication.

Helicase activation and origin unwinding are essential steps during DNA replication that expose ssDNA and thus have the potential to cause genomic instability. My studies have captured origin chromatin dynamics caused by an active helicase unwinding DNA, and have contributed evidence that the helicase may be intrinsically less processive in the absence of leading strand synthesis. These results may have implications for the mechanisms underlying human diseases involving polymerase α, and contribute to our growing understanding of how the eukaryotic cell preserves the integrity of the genome.

The complete and faithful duplication of the genome is essential to ensure normal cell division and organismal development. Eukaryotic DNA replication is initiated at multiple sites termed origins of replication that are activated at different time through S phase. The replication timing program is regulated by the S-phase checkpoint, which signals and repairs replicative stress. Eukaryotic DNA is packaged with histones into chromatin, thus DNA-templated processes including replication are modulated by the local chromatin environment such as post-translational modifications (PTMs) of histones.

One such epigenetic mark, methylation of lysine 20 on histone H4 (H4K20), has been linked to chromatin compaction, transcription, DNA repair and DNA replication. H4K20 can be mono-, di- and tri-methylated. Monomethylation of H4K20 (H4K20me1) is mediated by the cell cycle-regulated histone methyltransferase PR-Set7 and subsequent di-/tri- methylation is catalyzed by Suv4-20. Prior studies have shown that PR-Set7 depletion in mammalian cells results in defective S phase progression and the accumulation of DNA damage, which may be partially attributed to defects in origin selection and activation. Meanwhile, overexpression of mammalian PR-Set7 recruits components of pre-Replication Complex (pre-RC) onto chromatin and licenses replication origins for re-replication. However, these studies were limited to only a handful of mammalian origins, and it remains unclear how PR-Set7 impacts the replication program on a genomic scale. Finally, the methylation substrates of PR-Set7 include both histone (H4K20) and non-histone targets, therefore it is necessary to directly test the role of H4K20 methylation in PR-Set7 regulated phenotypes.

I employed genetic, cytological, and genomic approaches to better understand the role of H4K20 methylation in regulating DNA replication and genome stability in Drosophila melanogaster cells. Depletion of Drosophila PR-Set7 by RNAi in cultured Kc167 cells led to an ATR-dependent cell cycle arrest with near 4N DNA content and the accumulation of DNA damage, indicating a defect in completing S phase. The cells were arrested at the second S phase following PR-Set7 downregulation, suggesting that it was an epigenetic effect that coupled to the dilution of histone modification over multiple cell cycles. To directly test the role of H4K20 methylation in regulating genome integrity, I collaborated with the Duronio Lab and observed spontaneous DNA damage on the imaginal wing discs of third instar mutant larvae that had an alanine substitution on H4K20 (H4K20A) thus unable to be methylated, confirming that H4K20 is a bona fide target of PR-Set7 in maintaining genome integrity.

One possible source of DNA damage due to loss of PR-Set7 is reduced origin activity. I used BrdU-seq to profile the genome-wide origin activation pattern. However, I found that deregulation of H4K20 methylation states by manipulating the H4K20 methyltransferases PR-Set7 and Suv4-20 had no impact on origin activation throughout the genome. I then mapped the genomic distribution of DNA damage upon PR-Set7 depletion. Surprisingly, ChIP-seq of the DNA damage marker γ-H2A.v located the DNA damage to late replicating euchromatic regions of the Drosophila genome, and the strength of γ-H2A.v signal was uniformly distributed and spanned the entire late replication domain, implying stochastic replication fork collapse within late replicating regions. Together these data suggest that PR-Set7-mediated monomethylation of H4K20 is critical for maintaining the genomic integrity of late replicating domains, presumably via stabilization of late replicating forks.

In addition to investigating the function of H4K20me, I also used immunofluorescence to characterize the cell cycle regulated chromatin loading of Mcm2-7 complex, the DNA helicase that licenses replication origins, using H4K20me1 level as a proxy for cell cycle stages. In parallel with chromatin spindown data by Powell et al. (Powell et al. 2015), we showed a continuous loading of Mcm2-7 during G1 and a progressive removal from chromatin through S phase.

Abstract

DNA replication is an integral part of the cell cycle. Every time a cell divides, the entire genome has to be copied once and only once in a timely manner. In order to accomplish this, DNA replication begins at many points throughout the genome. These start sites are called origins of replication, and they are initiated in a temporal manner throughout S phase. How these origins are selected and regulated is poorly understood. Saccharomyces cerevisiae and Schizosaccharomyces pombe have autonomously replicating sequences (ARS) that can replicate plasmids extrachromosomally and function as origins in the genome. Metazoans, however, have shown no evidence of ARS activity.

DNA replication is a multistep process with several opportunities for regulation. Potential origins are marked with the origin recognition complex (ORC), a six subunit complex. In S. cerevisiae, ORC binds to the ARS consensus sequence (ACS), but no sequence specificity is seen in S. pombe or in metazoans. Therefore, factors other than sequence play a role in origin selection.

In G1, the pre-replicative (pre-RC) complex assembles at potential origins. This involves the recruitment of Cdc6 and Cdt1 to ORC, which then recruits MCM2-7 to the origin. In S phase, a subset of these pre-RC marked origins are initiated for replication. These origins are not fired simultaneously; instead, origins are fired in a temporal manner, with some firing early, some firing late, and some not firing at all.

The temporal firing of origins leads to wide regions of the genome being copied at different times during S phase. , which makes up the replication timing profile of the genome. These regions are not random, and several correlations between replication timing and both transcriptional activity and chromosomal landscape. Regions of the genome with high transcriptional activity tend to replicate earlier in S phase, and it is well know that the gene rich euchromatin replicates earlier than the gene poor heterochromatin. Additionally, areas of the genome with activating chromatin marks also replicate earlier than regions with repressive marks. Though many correlations have been observed, no single mark or transcriptional player has been shown to directly influence replication timing.

We mapped the replication timing profiles of three cell lines derived from Drosophila melanogaster by pulsing cells with the nucleotide analog bromodeoxyuridine (BrdU), enriching for actively replicating DNA labeled with BrdU, sequencing with high throughput sequencing and mapping the sequences back to the genome. We found that the X chromosome of the male cell lines replicated earlier than the X chromosome in the female cell line or the autosomes. We were then able to compare the replication timing profiles to data sets for chromatin marks acquired through the modENCODE (model organism Encyclopedia Of DNA Elements). We found that the early replicating regions of the male X chromosomes correlates with acetylation of lysine 16 on histone 4 (H4K16).

Hyperacetylation of H4K16 on the X chromosome in males is a consequence of dosage compensation in D. melanogaster. Like many organisms, D. melanogaster females have two X chromosomes while males have one. To compensate for this difference, males upregulate the genes on the X chromosome two-fold. This upregulation is regulated by the dosage compensation complex (DCC), which is restricted to the X chromosome. This complex includes a histone acetyl transferase, MOF, which acetylates H4K16. This hyperacetylation allows for increased transcription of the X chromosome.

We hypothesized that the activities of the DCC and the hyperacetylation of H4K16 also influences DNA replication timing. To test this, I knocked down components of the DCC (MSL2 and MOF) using RNAi. Cells were arrested in early S phase with hydroxyurea, released, and pulsed with the nucleotide analog EdU. The cells were arrested in metaphase and labeled for H4K16 acetylation and EdU. We found that male cells were preferentially labeled with EdU on the X chromosome, which corresponded with H4k16 acetylation. When the DCC was knocked down, H4K16 acetylation was lost along with preferential EdU labeling on the X chromosome. These results suggest that the DCC and H4K16 acetylation are necessary for early replication of the X chromosome. Additionally, early origin mapping of different cell lines showed that while ORC density does not differ between male and female cell lines, early origin usage is increased on the X chromosome of males, suggesting that this phenomenon is regulated at the level of activation, not pre-RC formation. Other experiments in female cell lines have been unclear about whether the DCC and subsequent H4K16Ac is sufficient for early X replication. However, these results are exciting because this is, to our knowledge, the first mark that has been found to directly influence replication timing.

In addition to these timing studies, I attempted to design a new way to map origins. A consequence of unidirectional replication with bidirectional replication fork movement is Okazaki fragments. These are short nascent strands on the lagging strand of replicating DNA. Because these fragments are small, we can isolate them by size and map them back to the genome. Okazaki density could tell us about origin usage and any directional preferences of origins. The process proved to be tedious, and although they mapped back with a higher density around ORC binding sites than randomly sheared DNA, little information about origin usage was garnered from the data. Additionally, the process proved difficult to repeat.

In these studies, we examined the replication timing program in D. melanogaster. We found that the male X chromosome replicates earlier in S phase, and this early replication is regulated by the DCC. However, it is unclear if the change in chromatin landscape directly influences replication or if the replication program is responding to other dosage compensation cues on the X chromosome. Regardless, we have found one the first conditions in which a mark directly influences the DNA replication timing program. 

All DNA-templated events, including replication and gene transcription, occur in the context of the local chromatin environment. The passage of the replication machinery results in disassembly of chromatin, which must be re-assembled behind the replication fork to re-establish the epigenetic state of the cell. Many of the factors and mechanisms regulating DNA replication and chromatin assembly have been identified from elegant in vitro biochemical experiments, work in model systems like Saccharomyces cerevisiae, or novel proteomic approaches. In spite of current advances in the field, it is still not clear how the chromatin landscape is organized and re-assembled during this process.

Current methods, while informative, lack the genome-wide base-pair resolution required to assess the dynamics of chromatin assembly and maturation in a spatial-temporal manner. To overcome the limitations of these studies, I have taken advantage of an epigenome mapping technique based on micrococcal nuclease (MNase) digestion followed by paired-end sequencing. This approach facilitates the analysis of chromatin structure by capturing not only nucleosomes, but also smaller DNA binding protein footprints in a factor-agnostic manner. I have developed a technique based on this approach that generates Nascent Chromatin Occupancy Profiles (NCOPs) to study the dynamics of chromatin assembly following passage of the DNA replication fork at a genome-wide level and at single base-pair resolution in S. cerevisiae. It employs a nucleoside analog to specifically enrich for nascent chromatin, which can be captured following a chase over different periods of time. Thus, NCOPs resolve the structure of nascent and mature chromatin, facilitating the analysis of chromatin maturation across the entire genome.

Using NCOPs, I provide a comprehensive description of the maturation process across different genomic regions and the dynamics of small DNA binding factor association with nascent and mature chromatin states. Our results support previous work characterizing the structure of nascent chromatin as being more disorganized and having poorly positioned nucleosomes. Importantly, using positioning and occupancy scores, I provide new details on the structure of nascent and mature chromatin at intergenic regions, including replication origins, and at highly transcribed and poorly transcribed genes. I uncovered that local epigenetic footprints have the potential to shape the dynamics of chromatin assembly, generating a chromatin maturation landscape that is dependent on the parental chromatin. Finally, I resolved patterns of transcription factor occupancy with nascent and mature chromatin, and observed transient factor association in the nascent state.

In all, this work provides insight into the dynamics of chromatin assembly, and allows for genome-wide and base-pair resolution investigation of chromatin maturation. The genomic and bioinformatic approaches developed here open the door for further investigation of the dynamics of epigenetic inheritance and the role of known and unknown players in re-establishing the eukaryotic epigenome following passage of the DNA replication fork.

Eukaryotic genomes have extensive flexibility and plasticity to modify transcription and replication programs, yielding a myriad of differentiated cell types and survival mechanisms to adverse environmental conditions. As these genomic processes require precise localization of DNA-binding factors, their dynamic temporal and spatial distributions provide dramatically different interpretations of a static genome sequence. DNA-binding factors must compete with nucleosomes, the basic subunit of chromatin, for access to the underlying DNA sequence. Even though the spatial preferences of these proteins are partially explained by DNA sequence alone, the complete genome occupancy profile has remained elusive, and we currently have a limited understanding of how DNA-binding protein configurations directly impact transcription and replication function.

Profiling the entire chromatin environment has typically required multiple experiments to capture both DNA-binding factors and nucleosomes. Here, we have extended the traditional micrococcal nuclease (MNase) digestion assay to simultaneously resolve both nucleosomes and smaller DNA-binding footprints in Saccharomyces cerevisiae. Visualization of protected DNA fragments revealed a nucleotide-resolution view of the chromatin architecture at individual genomic loci. We show that different MNase digestion times can capture nucleosomes partially unwrapped or complexed with chromatin remodelers. Stereotypical DNA-binding footprints are evident across all promoters, even at low-transcribed and silent genes. By aggregating the chromatin profiles across transcription-factor--binding sites, we precisely resolve protein footprints, yielding in vivo insights into protein-DNA interactions. Together, our MNase method, in one experiment, provides an unprecedented assessment of the entire chromatin structure genome-wide.

We utilized this approach to interrogate how the replication program is regulated by the chromatin environment surrounding DNA replication initiation sites. Pre-replicative complex (pre-RC) formation commences with recruitment of the origin recognition complex (ORC) to specific locations in the genome, termed replication origins. Although successful pre-RC assembly primes each site for S-phase initiation by loading the Mcm2-7 helicase, replication origins have substantially different activation times and efficiencies. We posited that replication origin function is substantially impacted by the local chromatin environment. Here, we resolved a high-resolution ORC-dependent footprint at 269 replication origins genome-wide. Even though ORC in S. cerevisiae remains bound at replication origins throughout the cell cycle, we detected a subset of inefficient origins that did not yield a footprint until G1, suggesting a more transient ORC interaction prior to pre-RC assembly. Nucleosome movement accommodated the pre-RC-induced expansion of the ORC-dependent footprint in G1, leading to increased activation efficiency. Mcm2-7 loading is preferentially directed to one side of each replication origin, in close proximity to the origin-flanking nucleosome. Our data demonstrates that pre-RC components are assembled into multiple configurations in vivo.

We anticipate that extending chromatin occupancy profiling to many different cell types will reveal further insights into genome regulation.

The DNA double-strand break (DSB) is one of the most toxic genomic lesions that can occur in any living cell. Failure to repair DSBs results in cell cycle arrest and ultimately programmed cell death, while improper repair can lead to profound alterations or loss of genomic information through translocations, inversions, deletions and other genomic aberrations. Although the molecular events required for the repair of double-strand breaks (DSB) have been well characterized, the role of epigenetic processes in the recognition and repair of DSBs has only been investigated at low resolution. I tested several site-specific DSB induction systems and found that that the HO endonuclease was able to rapidly and synchronously induce a site-specific DSB in Saccharomyces cerevisiae upstream of the PHO5 locus. This region of the genome is recognized for its chromatin organization, which is comprised of well-positioned nucleosomes. Utilizing MNase digestion of chromatin followed by paired-end fragment sequencing I was able to interrogate the order of chromatin changes that occur immediately following a DSB by generating a base-pair resolution map of the chromatin landscape. In wild-type cells, the first nucleosome left of the break was rapidly evicted. The eviction of this flanking nucleosome was dynamic and proceeded through an early intermediate chromatin structure where the nucleosome was repositioned in the adjacent linker DNA. Other nucleosomes bordering the break were also shifted away from the break; however, their loss was more gradual. These local changes preceded a broader loss of chromatin organization and nucleosome eviction that was marked by increased MNase sensitivity in the regions ~8 kb on each side of the break. While the broad loss of chromatin organization was dependent on the end-processing complex, Mre11-Rad50-Xrs2 (MRX), the early remodeling and repositioning of the nucleosome adjacent to the break was independent of the MRX and yKU70/80 complexes. I also examined the temporal dynamics of non-homologous end joining (NHEJ) mediated repair in a G1-arrested population. Concomitant with DSB repair, I observed the re-deposition and precise re-positioning of nucleosomes at the originally occupied positions. This re-establishment of the pre-lesion chromatin landscape suggests that a DNA replication-independent mechanism exists in G1 cells to preserve epigenome organization following DSB repair.

In eukaryotic cells, The local chromatin structures play a critical role in regulating all DNA-mediated events. However, the process of DNA replication is highly disruptive to the chromatin structure. Specifically, the parental chromatin needs to be disassembled before the passing of the replication fork and re-assembled afterward. Additionally, a maturation process is required for newly assembled chromatin to completely recover to the pre-replicative state. Rapid and faithful assembly and maturation of chromatin are essential for preserving the epigenome and maintaining genome integrity. Elegant in vitro biochemistry studies have allowed the characterization of numerous factors in the process of replication-coupled chromatin assembly. Recent advancements in genomic technologies further expanded our understanding of the process, providing genome-wide views of the spatial-temporal dynamics of chromatin maturation. However, there is still a lack of mechanistic details on many aspects of chromatin assembly, and the broader phenotypic significance of this process also remains unstudied.

One such example is Chromatin Assembly Factor 1 (CAF-1). It is one of the first histone chaperones identified that deposit histones onto the newly replicated DNA, yet there remain missing puzzles of CAF-1 ranging from the mechanisms by which CAF-1 interacts with DNA and the replisomes to its impact on chromatin maturation and how that relates to genome stability. Using nascent chromatin occupancy profiling (NCOP), I tracked the chromatin maturation kinetics in WT and CAF-1 mutant cells with high spatiotemporal resolution. I found that loss of CAF-1 results in a heterogeneous rate of nucleosome assembly, with individual nucleosomes exhibiting either rapid or slow maturation kinetics, ultimately leading to a global delay in chromatin maturation. The slow-to-mature nucleosomes are enriched with poly(dA:dT) sequences, suggesting CAF-1 facilitates (H3-H4)2 tetramer deposition and nucleosome maturation on sequences resistant to nucleosome formation. The assembly defect caused by loss of CAF-1 leads to the accumulation of nucleosome intermediates whose position is also influenced by underlying DNA sequences. The finding offers a new perspective on the pathway for nucleosome assembly in vivo. In addition, there exists a long-standing paradox that CAF-1 mutant cells exhibit loss of gene silencing and increased cryptic transcription, yet their steady-state chromatin landscape is nearly indistinguishable from that of WT cells. Our work illustrates that the dysregulation of transcription is temporary and occurs specifically in S phase, presumably as a result of transient defects in chromatin maturation. These results demonstrate how the DNA replication program can directly shape the chromatin landscape and regulate gene expression through the process of chromatin maturation.

During replication, the leading and lagging strands are replicated via distinct mechanisms. Numerous studies showed that different histone chaperones are employed for nucleosome assembly on the two strands; however, it remains unclear if or how the kinetics of chromatin maturation differ between the two strands. The current NCOP method does not distinguish actions on the leading and lagging strands. To address this, I developed strand-specific NCOP that captures the kinetics of chromatin maturation on the two daughter strands separately. Strand-specific NCOP experiments on CAF-1 and Mcm2 mutant cells provide evidence for the coordinated nucleosome assembly by Mcm2 and CAF-1 to ensure symmetric histone segregation and assembly on the daughter strands. Together, this work provides mechanistic insights into the role of multiple histone chaperones in chromatin assembly and maturation and opens up new avenues for understanding how disruption to these processes might contribute to defects in gene expression programs commonly found in disease states.

To ensure genomic integrity, the genome must be accurately duplicated once and only once per cell division. DNA replication is tightly regulated by replication licensing mechanisms which ensure that origins only initiate replication once per cell cycle. Disruption of replication licensing mechanisms may lead to re-replication and genomic instability.

DNA licensing involves two steps including the assembly of the pre-replicative compelx at origins in G1 and the activation of pre-RC in S-phase. Cdt1, also known as Double-parked (Dup) in Drosophila Menalogaster , is a key regulator of the assembly of pre-RC and its activity is strictly limited to G1 by multiple mechanisms including Cul4Ddb1 mediated proteolysis and inhibitory binding by geminin. Previous studies have indicated that when the balance between Cdt1 and geminin is disrupted, re-replication occurs but the genome is only partially re-replicated. The exact sequences that are re-replicated and the mechanisms contributing to partial re-replication are unknown. To address these two questions, I assayed the genomic consequences of deregulating the replication licensing mechanisms by either RNAi depletion of geminin or Dup over-expression in cultured Drosophila Kc167 cells. In agreement with previously reported re-replication studies, I found that not all sequences were sensitive to geminin depletion or Dup over-expression. Microarray analysis and quantitative PCR revealed that heterochromatic sequences were preferentially re-replicated when Dup was deregulated either by geminin depletion or Dup over-expression. The preferential re-activation of heterochromatic replication origins was unexpected because these origins are typically the last sequences to be duplicated during normal S-phase.

In the case of geminin depletion, immunofluorescence studies indicated that the re-replication of heterochromatin was regulated not at the level of pre-RC activation, but rather due to the restricted formation of the pre-RC to the heterochromatin. Unlike the global assembly of the pre-RC that occurs throughout the genome in G1, in the absence of geminin, limited pre-RC assembly was restricted to the heterochromatin. Elevated cyclin A-CDK activity during S-phase could be one mechanism that prevents pre-RC reassembly at euchromatin when geminin is absent. These results suggest that there are chromatin and cell cycle specific controls that regulate the re-assembly of the pre-RC outside of G1.

In contrast to the specific re-replication of heterochromatin when geminin is absent, re-replication induced by Dup over-expression is not restricted to heterochromatin but rather includes re-activation of origins throughout the genome, although there is a slight preference for heterochromatin when re-replication is initiated. Surprisingly, Dup over-expression in G2 arrested cells result in a complete endoreduplication. In contrast to the ordered replication of euchromatin and heterochromatin during early and late S-phase respectively, endoreduplication induced by Dup over-expression does not exhibit any temporal order of replication initiation from these two types of chromatin, suggesting replication timing program may be uncoupled from local chromatin environment. Taken together, these findings suggest that the maintenance of proper levels of Dup protein is critical for genome integrity.

Dim light vision requires the dopamine D1 receptor for the sensitization of rod bipolar cells by GABA. However, the circuit behind this mechanism is poorly defined. We used electroretinogram recordings of mice with genetic and pharmacological manipulations to investigate the identity of the cells in the circuit controlling rod bipolar cell sensitivity. We found that tonic GABA input onto rod bipolar cells comes from a wide-field amacrine cell rather than horizontal cells. The output of this wide-field amacrine cell is directly regulated by voltage-gated sodium channels and dopamine-dampened serial inhibition, providing multiple regulatory sites for adaptation in the rod system.

DNA replication is the process of synthesizing an exact copy of a genome during S-phase. DNA replication must only occur once and only once in the cell cycle. Therefore, the DNA replication program is a highly regulated process that is established prior to S-phase. In G1, the protein complexes that define an origin of replication are assembled stepwise onto chromatin. Potential origins of replication are first bound by ORC, origin recognition complex. ORC then recruits other factors that load the Mcm2-7 helicase onto the chromatin to assemble a pre-RC, pre-replication complex. Paradoxically, there is a vast excess of Mcm2-7 relative to ORC assembled onto chromatin in G1. These excess Mcm2-7 are broadly distributed on chromatin, exhibit little co-localization with ORC or replication foci, and can function as dormant origins. I used biochemical and genomic approaches to dissect the mechanisms regulating the assembly and distribution of the Mcm2-7 complex in Drosophila tissue culture cells. I found that Mcm2-7 loading occurs in two distinct phases during G1. In the first phase, limiting amounts of Mcm2-7 are loaded at ORC binding sites in a cyclin E/Cdk2 independent manner. Subsequently, there is a cyclin E/Cdk2 kinase activity dependent phase of Mcm2-7 loading that results in a 15-fold increase in chromatin associated Mcm2-7 and a dramatic genome-wide reorganization of the distribution of Mcm2-7 that is shaped by active transcription. Thus, increasing cyclin E/Cdk2 kinase activity over the course of G1 is not only critical for Mcm2-7 loading, but also the distribution of the Mcm2-7 helicase prior to S-phase entry.

The assembly of the pre-RC is not only required for DNA replication, but it has been implicated in being required for cohesin loading. The cohesin complex imparts cohesion between sister chromatids as they are replicated and remains in place until the sister chromatids are separated in mitosis. I assessed if pre-RC assembly is required for cohesin loading using genomic and biochemical approaches in Drosophila tissue culture cells. I found that pre-RC components co-localize with cohesin subunits throughout the Drosophila genome. I was unable to detect any cohesin loading onto chromatin mediated by pre-RC assembly or components in vivo. However, this result does not mean that they are not coordinated.

Any errors during DNA replication can cause genomic instability through rereplication, fragile sites, or stalled forks. In addition, other processes like sister chromatid cohesion that are coordinated with DNA replication can also introduce genomic instability. Aneuploidy is a potential consequence of sister chromatid cohesion defects resulting in unequal multiples of a genome within a cell. Aneuploidy can be detrimental to a cell or organism due to copy number variation (CNV) causing differences in expression of genes. However, cells are able to compensate for CNV between the sexes due to the differences in the number of sex chromosomes. I used genomic approaches to characterize three aneuploid Drosophila cell lines for the modENCODE project. I further characterized the S2 Drosophila cell line using immunofluorescence microcopy approaches to identify the chromosomal rearrangements that were mapped by de novo assembly of the genome. Both approaches showed that the S2 cell line has highly rearranged chromosomes. The S2 cell line was also analyzed to address if cells can compensate for CNV on autosomes using genomic approaches. In collaboration with the Oliver group (NIH) we found that S2 cells are able to compensate for CNV of autosomal genes by buffering gene expression.

In summary, my research explored mechanisms that a cell can employ to maintain genomic stability: assembly of dormant origins, chromosome segregation, and CNV compensation.

Advances in anti-cancer therapies, including chemotherapies, targeted therapies, and immuno-therapies, have drastically improved patient outcomes over the last few decades. However, tumor recurrence and acquired drug resistance continue to be detrimental for cancer patients, accounting for the majority of cancer-related deaths. Drug resistance is a common phenomenon that occurs when pharmaceutical agents are tolerated by and no longer effective against their target. In the case of cancer, acquired drug resistance is when cancer cells are able to survive, actively proliferate, and migrate to distant sites in the presence of anti-cancer agents. As such, it is imperative that we understand the mechanisms that lead to acquired drug resistance and tumor recurrence and leverage this newfound understanding to improve treatment options and, ultimately, patient outcomes.

Recurrent tumors and drug resistant cancer cells arise from drug-tolerant persister cells (DTPs), a population of cells that is able to withstand and adapt to cancer treatment. APOBEC3 and NRF2 are proteins that are largely absent and inactive in primary non-small cell lung cancer and HER2+ breast cancer, respectively. Interestingly, both proteins are found in recurrent disease, specifically after targeted therapy. This suggests that the upregulation of APOBEC3 and NRF2 is an adaptive response to anti-cancer treatment. Further, this suggests that APOBEC3 and NRF2 are essential for drug-tolerant persister cell survival and subsequent recurrence. Understanding how these proteins are induced, regulated, and what their respective roles are in tumor evolution can provide insight into therapeutic potentials in both residual and recurrent disease. First, I explored the regulation and function of APOBEC mutagenesis during acquired resistance to epidermal growth factor receptor (EGFR) inhibitors in two non-small cell lung cancer cell lines, PC9 and HCC827. I assess the effects of EGFR inhibition on APOBEC3 expression and activity and find that this induces APOBEC3 activity. Moreover, I evaluate how sustained APOBEC3 activity promotes evolution of drug resistance in DTPs. Finally, I examine what mechanisms might play a role in acquired drug resistance when APOBEC3 is highly active. I show that while APOBEC activity does not accelerate acquired therapy resistance, A3B expression alters the evolutionary path that PC9 cells take to become gefitinib-resistant. Specifically, A3B expression is associated with the late acquisition of T790M mutations during the DTP state, supporting a model where induction of APOBEC activity promotes DTP survival, thereby facilitating the on-going evolution of drug-tolerant persister cells. Of note, I find that APOBEC3 activity is associated with squamous cell transdifferentiation in PC9 cells, suggesting that p63 and its target genes could be future biomarkers and therapeutic targets.

Second, I investigated the regulation of the transcription factor NRF2 in recurrent breast cancer cells. I assess KEAP1-mediated regulation of NRF2 and find that while KEAP1 knockout in primary and recurrent cells caused an increase in NRF2 and its target genes, NRF2 remained more elevated in recurrent cells, indicating that increased NRF2 levels and transcriptional activity in these cells are independent of KEAP1. Next, I looked at post-translational modifications on NRF2 to determine if this may be the cause of the differential NRF2 levels in primary and recurrent cells. I found that NRF2 in recurrent cells had higher levels of phospho-Ser364, potentially affecting NRF2’s stability. Finally, I evaluated regulation of NRF2 by Akt and GSK-3β. I found that inhibition of Akt had no effect on NRF2 levels. In contrast to this, I found that GSK-3β activity is inversely correlated with NRF2 levels, suggesting that low levels of GSK-3β activity is partially responsible for NRF2 stabilization in recurrent tumor cells.

In conclusion, I have modeled two adaptive mechanisms for tumor recurrence and acquired drug resistance in two different cancer types. I elucidated the mechanisms by which APOBEC3B and NRF2 are able to promote cancer cell evolution in drug-tolerant persister cells that eventually give rise to recurrences.