We propose to develop a nanopore-based nanofluidics platform for detection and analysis of single protein molecules from live cells. This platform would consist of two parts – first a microfluidic system to capture cells and generate micro-ruptures or electropores in their membranes, so that proteins can be extracted, and second a nanogap-nanopore device, consisting of a nanopore ~1 to 10 nm in diameter between metal electrodes on a silicon nitride (SiN) membrane substrate, to analyze the extracted molecules. Nanogap-nanopore devices are made by nanosculpting metal films using a transmission electron beam. Recently, we have demonstrated the fabrication of metal nanostructures and integrated devices of arbitrary shape with unprecedented resolution down to sub-nm precision using transmission electron beam ablation lithography (TEBAL). Compared to the existing nanopore-based molecular analysis configuration described below, these devices add significant new sensing capabilities. Transverse electrodes placed directly adjacent to the nanopore allow sensing of the conduction and capacitance of just the nanometer-sized gap between the electrodes, and the resulting changes when molecules translocate through the nanopore. Similarly, charged regions of a molecule will induce an opposite charge on the electrodes, which can be measured as a voltage signal.
This combination of nanopore-based analysis with a method for extracting protein molecules from live cells is a powerful new technology for real-time analysis of biological processes in live cells.
Cellular responses to DNA damage are critically important for both suppressing and treating malignancy. There is substantial evidence that nearly all cancers experience elevated DNA double strand breaks (DSBs) at early stages of development and that therapeutic responses to anti-cancer chemotherapy are tightly linked to repair efficiency. The strength of the DSB repair response thus appears to be a pivotal determinant of cancer susceptibility and chemotherapeutic response. The breast cancer susceptibility protein, BRCA1, recognizes lysine63-linked ubiquitin chains (K63-Ub) at DSBs via a recently reported interaction with the Rap80 protein. Rap80 contains tandem ubiquitin interaction motifs (UIMs) that specifically recognize K63-Ub and are both necessary and sufficient for DSB localization. Ubiquitin chain synthesis is initiated within minutes of DSB induction by an ATM kinase (Ataxia Telangiectasia Mutated) dependent phosphorylation cascade and both phosphorylation and ubiquitin removal coincides with the conclusion of DSB repair. Thus, phosphorylation and ubiquitin chain dynamics at DSBs tightly mirror the DNA damage response and are accurate indicators of repair efficacy. Currently, no methods are available to monitor DNA damage specific ubiquitin chain dynamics in vivo. Studies have instead relied on examination of fixed cells and tissues, precluding an evaluation of DNA damage response/checkpoint activation dynamics in the context of the “lifetime” of a tumor, (i.e. during carcinogenesis and responses to chemotherapy). An in vivo method to interrogate DSB accumulation and repair during (1) tumor development and (2) therapeutic responses; would thus provide an invaluable resource. The goal of this proposal is to develop an in vivo system to analyze DSB repair activation dynamics in individual cells by exploiting the specificity of the Rap80 UIM domains to detect ubiquitin chain dynamics at DSBs. This system would represent the first of its kind in terms of monitoring both the DSB repair response and in sensing real time ubiquitination events.
RNA molecules fold into three-dimensional structures through specific base-pairing interactions that are encoded within their sequences. Recent discoveries have revealed RNA molecules themselves perform a variety of tasks, from gene expression regulation to catalytic activities, and this functionality is intimately linked to their three-dimensional structures. Furthermore, the functionality of the highly conserved RNA silencing pathway is completely dependent on the three-dimensional structures of RNA molecules. Specifically, long double-stranded RNAs (dsRNAs) or self-complementary RNA fold-back structures give rise to small RNAs (smRNAs) through the activity of RNase III-type ribonucleases, and these smRNAs comprise the sequence-specific effectors of RNA silencing pathways that direct the negative regulation or control of genes, repetitive sequences, viruses, and mobile elements. Thus, it is not surprising that a growing number of diseases have been tied to the deregulation of RNA expression, processing, as well as formation of its secondary structure. However, the exact mechanisms by which these molecular defects drive disease progression are unknown. Therefore, the aim of this proposal is to develop new high-throughput methodologies focused on structural studies of RNA, which will allow a fundamental increase in our understanding of the roles of this molecule in genetic regulation, as well as human development and disease.
Compartmentalization of the cytoplasm into membrane-bound organelles is a fundamental feature of the evolution of prokaryotic cells to eukaryotic cells. Highly coordinated transport mechanisms direct intracellular molecules to their defined locations and ensure that the identity and function of individual compartments are maintained. Membrane trafficking in the secretory and endocytic pathways is a multistep process involving the generation of transport carriers (vesicles) loaded with defined sets of cargo, their shipment between compartments, and their specific fusion with the target membranes. These membrane-mediated processes involve a complex array of protein and lipid interactions and are tightly regulated by intracellular signaling events, in which a large role belongs to posttranslational modifications. This regulation has been extensively studied over the past decades, however the pathways that regulate membrane trafficking are still poorly understood. One of the technical challenges in the studies of intracellular membrane trafficking is continuous live imaging of vesicles at different stages of the transport pathways. Current imaging technologies enable live observations of intracellular vesicle movements and their responses to stimuli, or high resolution structural imaging of individual vesicles; however, there are currently no methods that allow tracking the intracellular fate of individual vesicles during their transitions between intracellular compartments. Our current project is focused on developing such a method, making use of pH changes that occur inside the vesicles during their inter-compartment trafficking. To do this, we are developing a new macromolecular fluorescent pH-sensitive probe that will be taken up by cells via receptor-mediated endocytosis and will undergo color changes indicative of the pH in its immediate environment (e.g. endosome vs lysosome). By using multichannel optical registration, two-fold information will be obtained: spatial localization of organelles containing the probe, and the pH in these organelles. We will validate this method by in vivo observation of vesicle trafficking during receptor-mediated endocytosis in cultured cells and comparing it with the known molecular markers of different intracellular compartments. We will further apply the method to study the regulation of vesicle and membrane trafficking by protein arginylation - a poorly understood posttranslational modification with an emerging role in the regulation of intracellular signaling events.
Real-time Proteomics (RTP) is a powerful technology to visualize the synthesis of individual proteins in live cells. This method has dramatic implications for both basic biomedical research and for the development of new diagnostics and therapies. RTP takes advantage of FRET-based methods to monitor real-time protein biosynthesis. In this technology, the specific binding of fluorescently labeled functional tRNAs to fluorescently labeled functional ribosomes produces a FRET signal. The timing, pattern, and duration of this FRET signal is a highly sensitive metric for translational rate, fidelity, and processivity by the ribosome. The ability to monitor tRNA-ribosome interactions in real time allows us to ‘watch’ protein translation at the single-molecule level. The major aims of this project are to implement RTP at the single molecule in vitro level and at the single cell ensemble level in live C. elegans. With RTP, we hope to be able to probe the role of the ribosome and protein translation in diverse biological processes in ways that are not currently possible due to lack of enabling technologies.
Protein kinases are critical for the regulation of many biological processes, and temporal and spatial control of phosphorylation underlie many cellular functions. Measurements of phosphorylation dynamics in living cells can provide a powerful representation of the state of a complex biological system in its native cellular context. We will examine these dynamics with high temporal and spatial resolution, using fluorescence-based sensors that report on the phosphorylation states of localized peptide substrates in living cells. This proposal is motivated by specific questions related to cell division and cancer development, but the technologies that we are developing will be generally applicable to a wide range of biological problems. These technologies include (1) design of phosphorylation sensors targeted to specific intracellular structures, (2) unbiased, high-throughput methods to screen libraries of such sensors, (3) image analysis methods to track phosphorylation dynamics in four dimensions (x, y, z, and time), and (4) development of zebrafish as a model system to examine phosphorylation dynamics in the context of a living vertebrate organism.
This proposal seeks to utilize primary human macrophages in cell-based screens to identify essential components of anti-microbial activity against the experimentally tractable, protozoan parasite Toxoplasma gondii. Macrophages are immune cells that exhibit potent killing activity against a broad range of pathogens. After 'sensing' parasites, viruses or bacteria via pathogen-associated molecular patterns, macrophages become activated by inflammatory signals and initiate transcriptional programs that typically result in cell-autonomous destruction of intracellular pathogens. The intrinsic ability of these cells to control a wide range of infectious agents makes macrophages an appealing cell type in which to conduct genome-wide functional screens. The results from our screens would identify human genes involved in innate immunity to intracellular pathogens and would be of immense interest to Toxoplasma researchers, as well as the broader community of cellular immunologists. In addition, once the basic methodology outlined above is optimized, it should be feasible to extend this screen to compare and contrast how macrophages restrict other important pathogens, including Mycobacterium tuberculosis and HIV.
Retroviruses exhibit a high degree of cell type and host range specificity that is in part determined by the cellular entry receptor. Several years ago, we identified mouse transferrin receptor (TfR1) as the mouse mammary tumor virus (MMTV) receptor. TfR1, a type II membrane glycoprotein, is the major means by which cells take up iron. The receptor has a high affinity for iron loaded transferrin (Tf) at neutral pH, and upon binding ligand, is endocytosed via clathrin coated pits to the early acidic endosome (pH6.0) where iron is released. After release, TfR1 recycles back to the cell surface, releases Tf and a new molecule of iron loaded Tf is bound. TfrR1 expression is regulated at both the transcriptional and posttranscriptional levels. As a result, when there is sufficient iron, TfR1 levels are low and conversely, when iron stores are low, receptor levels are high. TfR1 is also an activation marker on lymphoid cells and MMTV, which as a milk-borne virus first infects dendritic cells and then lymphoid cells in the small intestine, probably induces TfR1 expression to gain entry into the host. MMTV requires entry from a late (pH5.0) acidic compartment rather than the early acidic compartment to which TfR1 traffics upon Tf binding and although it enters cells bound to TfR1, infection is independent of clathrin mediated endocytosis. To determine how MMTV enters cells, we propose to use high throughput cell based screening to identify host cell molecules important for MMTV infection. This will parallel a screen just beginning in our lab to look at the intracellular requirements for infection of the New World arenavirus, Junin Virus (JUNV), which causes hemorrhagic fevers in humans. Like MMTV, JUNV uses TfR1 as its entry receptor, but enters the organism through aerosols i.e. the lungs and requires clathrin-mediated endocytosis. Interestingly, MMTV and JUNV use different regions of the mouse and human TfR1, respectively, to achieve infection. Thus, we believe that this parallel screen is likely to yield important data about the cell biology of virus entry and pathogenesis.
The goal of our project was to explore the interaction of the mammalian immune system with microbial communities associated with mammals, both during healthy mutualism and pathogenesis. In our first study, we investigated the spatial and temporal response of the murine gut microbiome to infection with Citrobacter rodentium, an attaching-effacing bacterium that provokes innate and adaptive immune responses, resulting in transient bacterial colitis. We used DNA bar coding and 454 pyrosequencing to characterize 102,398 partial 16S rDNA sequences from 85 microbial samples from tissue adhered and luminal bacteria of the cecum, proximal colon, and distal colon. The deep sequencing data revealed that C. rodentium was most abundantly associated with the cecal mucosa at day 9 post-infection, and then diminished in abundance, providing the first report of use of deep sequencing to track a pathogen in vivo through the course of infection. Notable changes were associated with both the mucosally-adhered and lumenal microbiota at both day 9 and day 14 p. i. Alterations in abundance were seen for Proteobacteria, Deferribacteres, Clostridia and others. The Lactobacillus group dropped in abundance during infection, which may be important for pathogenesis because members of this lineage modulate the composition of the gut microbiota and are used as probiotics. Thus deep sequencing provides previously inaccessible information on how Citrobacter infection and clearance reshapes the gut microbial community in space and time. In our second study, we used deep sequencing to characterize the murine microbiome in the presence of a high fat diet. We compared 25,790 16S rDNA sequences from both wild type and Relm-beta knockout mice on low fat and high fat diets. We found large alterations associated with switching to the high fat diet, including an increase in the Firmicutes, a decrease in Bacteroidetes, and an increase in Proteobacteria. This was seen for both genotypes (which corresponded to the presence and absence of obesity), indicating that the high fat diet itself, and not the obese state, mainly accounted for the observed changes in the gut microbiota. The RELM-beta genotype also independently influenced microbiome composition detectably. Metagenomic analysis of 537,604 sequence reads documented extensive changes in gene content due to a high fat diet, including an increase in transporters and two-component sensor-responders as well as a general decrease in metabolic genes. Unexpectedly, we found a substantial amount of murine DNA in our samples that increased in proportion on a high fat diet. These results demonstrate the importance of diet as a determinant of gut microbiome composition and suggest the need to adjust for dietary variation when analyzing the possible role of the gut microbiome in human disease.
Human-specific LINE-1 elements (L1Hs) are the only active family of LINE-1 retrotransposons in the human genome. Throughout evolutionary history there has been a succession of L1 families which through their copy-and-paste replication mechanism are directly responsible for ~17% of genomic sequence. The active L1Hs family is still inserting new copies into the genome as evidenced by ~250 polymorphic insertion sites catalogued in several studies and at least 18 known cases of a de novo L1 insertion contributing to human diseases. These known polymorphic and private insertions are likely to represent only a small fraction of the total amount of genomic variation induced by L1 activity. To address this possibility, we have developed an assay whereby the Human-specific LINE-1 content in the genome of a given individual can be assessed in a high-throughput manner, on a genome-wide scale. Following amplification of the 3’ flanking sequences via a PCR-based approach, amplicon ends are sequenced on the Illumina Genome Analyzer platform yielding millions of reads per individual. These reads are analyzed via a computational pipeline whereby clusters of reads indicate the presence of a human-specific L1 insertion which can then be validated by PCR. Our results indicate that in excess of 100 insertions are present in any given individual that are not represented in the reference human genome sequence. Studies using this technique to directly measure the rate of retrotransposition in humans and examine L1 allele frequencies at the population level are underway.
Missed the Sept. 30 lecture-discussion on genomics & stem cells? Watch the video here.
Basic Wet Lab Techniques for Genomics Research Workshop (for faculty), Nov. 13-20 - more info.
PGFI's Visiting Scholars program is soliciting nominations.