Duplication of the genome prior to cell division occurs in a spatially and temporally organized manner. The temporal order of replication (replication timing) reflects the higher order organization of the genome. During the first half of the S-phase, euchromatic regions are replicated followed by facultative heterochromatin during mid S-phase and finally constitutive heterochromatin regions in the second half of the S-phase. In the nucleus, euchromatin is localized in the interior while constitutive heterochromatin is rather located at the periphery. The spatio-temporal program of genome replication changes throughout development and cellular differentiation and has been correlated with changes in chromatin dynamics, histone marks, and nuclear architecture indicating that genome reorganization is associated with epigenetic and gene expression changes. Abnormal replication timing has also been reported in many diseases, including cancer. A growing body of evidence indicates that the replication-timing program strongly influences the spatial distribution of mutagenic events such that certain regions of the genome that are replicated in late S-phase present increased spontaneous mutagenesis compared to surrounding regions replicated in early S-phase. This has been observed during the evolution of species as well as during the evolution of cancer. In addition, a recent report showed that the dominant determinant of regional mutation rate variation is chromatin organization, with mutation rates elevated in more heterochromatin-like domains and repressed in more open chromatin. Although different hypotheses have been advanced, the cause of this increasing gradient of point mutation rates with later replication timing and the underlying mechanisms remain elusive. Although no mutagenic profile has been reported so far from pluripotent cells undergoing differentiation, one can expect that genome reorganization influences the mutagenic landscape. Obvious and important questions emerge from these observations: how and why elevated mutation rates are associated with heterochromatin-like domains which are replicated in late S-phase? What can be the “cost” for the cell when global genome reorganization takes place after differentiation/dedifferentiation (e.g. pre-cancerous cells) or reprogramming (e.g. iPS cells) if late-replicating regions containing point mutations become early and are actively transcribed? Point mutations arising in the genome are mainly caused by a special class of DNA polymerases (TLS polymerases) that support replication directly past template lesions (or unusual DNA secondary structures) that cannot be negotiated by the replicative high-fidelity polymerases. However, these specialized polymerases can be highly error-prone on undamaged DNA. An emerging concept proposes that these enzymes may also function during the unchallenged S-phase. On the basis of solid preliminary results, we assume that the essential error-prone DNA polymerase zeta (Pol zeta) is required to replicate through condensed chromatin regions and could be involved in the gradient of spontaneous mutagenesis. Therefore, this ambitious SPUR project aims to decipher the involvement of Pol zeta??and its catalytic subunit Rev3) in the regulation of the spatio-temporal program of DNA replication during embryonic stem cell differentiation and comprehensively evaluate the role of this error-prone polymerase in the point mutation frequencies which increase in late-replicating regions.

Due to their capacity to jump from one site to another, transposable elements (TEs) are a significant threat to genome integrity. To limit TE mobilization, most eukaryotes have evolved small RNAs (sRNAs) to silence TE activity via homology-dependent mechanisms. sRNAs guide PIWI proteins to which they bind to homologous TE sequences, recruit histone-modifying enzymes, and repress the transcriptional activity of TEs. However, how small RNA-PIWI complexes recruit downstream effectors remains elusive. We will address this important question using the sRNA-guided genome elimination paradigm in ciliates. In the ciliate Paramecium, massive and reproducible elimination of TEs occurs during the development of the somatic genome from the germline genome at each sexual cycle. The specific recognition of TEs involves sRNAs which direct heterochromatin formation and subsequent TE elimination. sRNAs are produced from the entire germline genome, from TEs and non-TE sequences, by a developmental-specific RNAi interference pathway. Non-TEs sRNAs, which represent a large fraction of the sRNA population, are degraded, while sRNAs corresponding to TEs will remain and recruit Polycomb Repressive Complex 2 (PRC2) and trigger TE elimination. How non-TEs sRNAs are massively degraded and TE sRNAs are selected is currently unknown. In this proposal, we aim at uncovering how sRNAs are selectively degraded in Paramecium, an exquisite model because of the abundance and precise developmental timing of sRNA degradation. Our preliminary data identified new factors required for the process. We will use multidisciplinary, cutting-edge approaches, combining functional genomics and proteomics, to decipher the mode of action of these factors and the pathways involved.

Precise control of the transition from self-renewal to terminal differentiation in stem cells is critical to maintain a balance between cell populations: an excess of stem cell self-renewal can lead to tumourigenesis, whereas an excess of differentiation can deplete the stem-cell pool. In the adult Caenorhabditis elegans germline, Notch signals emanate from the somatic distal tip cell to maintain germline stem cells (GSCs) in a proliferative state by repressing the translation of meiotic promoting factors. We have uncovered a novel pathway regulating the decision between GSC renewal and meiotic differentiation that involves the ubiquitin-proteolytic system (UPS). Using a novel temperature-sensitive allele of the cul-2 gene, we found that the CUL-2 RING E3 ubiquitin ligase, in combination with the Leucine Rich Repeat 1 substrate recognition subunit (CRL2LRR-1), negatively regulates the transition from the mitotic zone of the germline to the meiotic programme of chromosome pairing, synapsis, and recombination. More specifically, we find that CRL2LRR-1 regulates in stem cells the stability of the HORMA domain-containing protein HTP-3, which is required for loading structural proteins onto meiotic chromosomes and for the formation of the double-strand breaks that initiate meiotic recombination. Furthermore, we found that cyclin E/Cdk2 kinase, which is specifically activated in GSCs but repressed upon meiotic differentiation, phosphorylates HTP-3 and regulates its stability. Besides HTP-3, CUL-2 targets other factors for degradation to prevent precocious meiotic entry and to promote germline stem cell proliferation. Herein, we propose to use a unique combination of genetics, cell biology, biochemical and quantitative proteomic approaches to elucidate the role of protein degradation in germ cell biology. In particular, we propose to identify CUL-2 targets in the germline and the molecular mechanisms controlling their degradation in space. The role of the UPS and CUL-2 in germline stem cell biology has not been studied so far. CUL-2 is evolutionarily conserved in metazoans and appears to regulate germ cell divisions in Drosophila. Therefore emerging paradigms provided by the study of germ cell biology in C. elegans should be directly applicable in other systems and possibly also in humans.

One of the most challenging problems in modern biology is to address at the experimental level the analysis of the dynamics of proteome variations in composition and structure in order to decipher the fundamental mechanisms of gene expression and regulation in normal and pathological conditions. Using U-[12C]-glucose as the sole source of carbon to grow prototrophic cells, we developed a Simple Light Isotope Metabolic (SLIM) labeling strategy highly effective to analyze with an unprecedented depth complex proteomes in bottom-up and top-down experiments. We want now to elaborate an automatic workflow processing of MS bottom-up raw data for quantitative proteomics. This requires robust statistical analysis of the SLIM-labeling based quantitative proteomics procedures. We want to apply the SLIM-labeling strategy to analyze the quantitative variations of proteoforms in top-down experiments, addressing the complexity of proteome from multi-cellular organisms and higher eukaryote cells.

Formins are key regulators of actin assembly, which promote the rapid elongation of filaments, leading to typical elongated filament structures such as filopodia or the cytokinetic ring. Formins are involved in a number of important cell processes and related pathologies, such as angiogenesis, neuropathies, and cancer. Formins form homodimers that are able to track an elongating filament barbed end (using their FH2 domains) and to drastically enhance its elongation from profilin-actin (FH1 domains), resulting in the rapid formation of long filaments. Over the past ten years, formins have been scrutinized intensely and several of their key features have been unveiled. Strikingly, the conformational changes involved in formin activity, which are essential to understand how formins function, are still speculative. These functional conformations have never been observed. Rather, a model of formin activity has emerged, based on indirect biochemical, biophysical and structural data. For instance, it is commonly assumed that the FH2 dimer translocates at the growing barbed end, in rapid equilibrium between two conformations, and that the FH1 domain forms a flexible chain which can explore the space around the filament and deliver subunits to the growing end. Recent observations have given rise to a number of paradoxical results, showing that this model, while it may be broadly correct, fails to account for important features of formins. Today, our understanding of formin function is limited by our ignorance of the conformations it adopts in order to perform its amazing tasks. Here, we propose to combine different in vitro approaches to decipher formin’s functional conformations, and test some predictions arising from our preliminary data. Using different electron microscopy techniques, we will observe the conformations adopted by the FH2 dimer in the various situations it can encounter while interacting with the actin filament. Local modifications of the filament conformation will also be scrutinized. The affinity of the FH1 chain for profilin and profilin-actin will be measured, and it will be manipulated with magnetic tweezers in order to test its mechanical properties. The coupling between FH1 extension and the binding of profilin-actin monomers will be specifically investigated. Possible interactions between the FH1 and FH2 domains, which could explain some recent and surprising results, will also be addressed. The coordinator’s lab (partner 1) is an expert of actin assembly and has recently produced a number of important results on formins, using biochemistry and a novel microfluidics approach combined with optical microscopy. Partner 1 will thus be able to produce the proteins required for this project and generate the appropriate samples for electron microscopy observations, which will be carried out by partner 2 who is an expert of these techniques, as well for the magnetic tweezer experiments, which will be done by partner 3 who is a renowned expert of molecular manipulations. We are thus confident that our combined expertise will lead this ambitious project to success.