T cell dysfunction is a key problem in cancer, enabling not only tumorigenesis but also causing resistance to immunotherapy. Induction of immune checkpoints is a hallmark of T cell dysfunction, but clinical blockade by PD-1 and CTLA-4 antibodies solves this problem for few patients only. Dysfunction is driven also by additional mechanisms, including chronic stimulation and metabolic insufficiency. A better mechanistic understanding will be imperative for improving immunotherapy. My laboratory recently launched “ReverT”, a genome-wide CRISPR-Cas9 screening program to identify genes, ablation of which reverses dysfunction in primary T cells. To de-risk this application, we have already successfully completed three such screens in vitro, for chronic antigen stimulation, metabolic stress and PD-1 induction, validating and characterizing several dysfunction genes not previously reported. Furthermore, an independent follow-up re-screen with a Dysfunction Reversion Candidate (DRC) mini-library containing the top 100 hits of each of these screens validated >100 genes that reversed T cell dysfunction in vitro. Here, we will carry out the most challenging and important step: to systematically validate and mechanistically characterize this collection of T cell dysfunction genes in vivo. This will be done in a pooled and multimodal fashion, analyzing multiple dysfunction phenotypes in parallel, specifically immune checkpoints, exhaustion, metabolism, recruitment and proliferation. Our proof-of-concept results indicate that the DRC library contains “nodal” factors, operating in several seemingly different dysfunction settings, which may thus in fact be linked. We will use a collection of adoptive cell transfer mouse and human tumor models for validation and mechanistic characterization, as well as primary human T cells in patient-derived tumor fragments. Lastly, we will translate our findings to a preclinical setting, aiming to achieve more durable clinical responses.

When DNA is transcribed or replicated, torsional stress accumulates on the double helix. This tension must be dissipated by spreading it along the DNA fiber, or it must be removed altogether. One of the main factors responsible for the removal of such stress is DNA Topoisomerase I (Top1), an important target of cancer chemotherapy. When Top1 activity is lost, torsional stress accumulates on transcriptionally active genes and can lead to the formation of non-canonical DNA/RNA hybrid structures called R loops. These structures are emerging as important regulators of genome function and stability. By genome-wide mapping of R loops in human cells, I recently found that depletion of Top1 leads to a marked R loop stabilization, specifically on genes that are anchored to the nuclear lamina. This strongly suggests that attachment of DNA to the nuclear lamina may prevent dissipation of torsional stress, but how this works is still largely unclear. I propose to investigate the causal relationships between torsonal stress, Top1, R-loops and nuclear lamina attachment, taking advantage of a suite of unique genomics techniques developed in the host lab. Specifically, I will: 1) Develop two novel reporter assays to probe the effects of chromatin context (in particular lamina associated chromatin) and Top1 on torsional stress and R loop formation, at thousands of locations in the human genome. 2) Investigate if and how Top1 regulates DNA/nuclear lamina contacts by by means of a novel version of the powerful genome-wide DamID mapping method with much-improved time resolution. My expertise in Top1, R-loops and DNA topology combined with the unique genomics methodologies in the host lab, as well as their expertise in lamina-associated DNA, will lead to a unique synergy that should result in new insights into the relationship between nuclear organization, torsional stress and R loop formation. Moreover, it will yield new methods that will boost scientific progress in this field.

The 3D organization of chromosomes within the nucleus is of great importance to control gene expression. The cohesin complex plays a key role in such higher-order chromosome organization by looping together regulatory elements in cis. How these often megabase-sized looped structures are formed is one of the main open questions in chromosome biology. Cohesin is a ring-shaped complex that can entrap DNA inside its lumen. However, cohesin’s default behaviour is that it only transiently entraps and then releases DNA. Our recent findings indicate that chromosomes are structured through the processive enlargement of chromatin loops, and that the duration with which cohesin embraces DNA determines the degree to which loops are enlarged. The goal of this proposal is two-fold. First, we plan to investigate the mechanism by which chromatin loops are formed, and secondly we wish to dissect how looped structures are maintained. We will use a multi-disciplinary approach that includes refined genetic screens in haploid human cells, chromosome conformation capture techniques, the tracing in vivo of cohesin on individual DNA molecules, and visualization of chromosome organization by super-resolution imaging. With unbiased genetic screens, we have identified chromatin regulators involved in the formation of chromosomal loops. We will investigate how they drive loop formation, and also whether cohesin’s own enzymatic activity plays a role in the enlargement of loops. We will study whether and how these factors control the movement of cohesin along individual DNA molecules, and whether chromatin loops pass through cohesin rings during their formation. Ultimately, we plan to couple cohesin’s linear trajectory along chromatin to the 3D consequences for chromosomal architecture. Together our experiments will provide vital insight into how cohesin structures chromosomes.

The human genome carries genetic information in two distinct forms: Transcribed genes and regulatory DNA elements (rDEs). rDEs control the magnitude and pattern of gene expression, and are indispensable for organismal development and cellular homeostasis. Nevertheless, while large-scale functional genetic screens greatly advanced our knowledge in studying mammalian genes, such tools to study rDEs were lacking, impeding scientific progress. Interestingly, recent advance in genome editing technologies has not only expanded the available screening toolbox to examine genes, but also opened up novel opportunities in studying rDEs. We distinguish two types of rDEs: Transcriptional rDEs that recruit transcription factors to enhancers, and structural rDEs that maintain chromatin 3D structure to insulate transcriptional activities, a feature postulated to be essential for gene expression regulation by enhancers. Recently, we developed a CRISPR strategy to target enhancers. We showed its scalability and effectivity in identifying potential oncogenic and tumour-suppressive enhancers. Here, we will exploit this line of research and develop novel strategies to target structural rDEs (e.g. insulators). By setting up functional genetic screens, we will identify key players in cell proliferation, differentiation, and survival, which are related to cancer development, metastasis induction, and acquired therapy resistance. We will validate key insulators and decipher underlying mechanisms of action that control phenotypes. In a parallel approach, we will analyse whole genome sequencing datasets of cancer to identify and characterize genetic aberrations occurring in the identified regions. Altogether, the outlined research plan forms a natural extension of our successful functional approaches to study gene regulation. Our results will setup the foundation to better understand principles of chromatin architecture in gene expression regulation in development and cancer.