Cell cycle regulation of structure-selective endonucleases during homologous recombination

The eukaryotic cell cycle is a complex process that coordinates protein function with the changing requirements of the different cell cycle phases. Many proteins are therefore regulated in a cell cycle-specific manner to make them available/active at a specific cell cycle phase, or prevent their action at other phases. Two proteins regulated in such a cell cycle-specific manner are the structure-selective endonucleases (SSEs) Mus81-Mms4 and Yen1 – repair factors required for the removal of DNA structures arising during homologous recombination (HR). Research in the last years thereby identified a variety of regulatory pathways leading to cell cycle-specific upregulation of the Mus81-Mms4 and Yen1 catalytic activity during M-phase. Despite accumulating evidence that the catalytic activity of the two SSEs is cell cycle-regulated, it remained elusive at which cell cycle phase they would exhibit their key function and how the different regulatory mechanisms upregulating Mus81-Mms4 and Yen1 during M-phase are working together. To address these questions, we developed an advanced toolbox of cell cycle tags which allowed us to restrict the expression of Saccharomyces cerevisiae Mus81-Mms4 and Yen1 to different cell cycle phases and thus analyze at which cell cycle phase these SSEs exhibit their key function. The advanced toolbox of cell cycle tags generally refines the methodology of cell cycle tags and overcomes critical limitations observed for previous cell cycle tag systems, such as the limited number of cell cycle tag constructs that did not allow adaption of expression levels. We circumvented this problem using genetic approaches like chimeric protein fusions, 5´UTR truncations and out-of-frame ATGs which resulted in a toolbox of 46 cell cycle tag constructs with a broad range of expression levels. In general, these advancements will help to answer the question of cell cycle regulation for many proteins and, more specifically, allowed us to address this question for the SSEs Mus81-Mms4 and Yen1. Applying the advanced cell cycle tag toolbox to Mus81-Mms4 and Yen1, we were able to restrict their expression to different cell cycle phases and attribute their key function to M-phase. Furthermore, we used the approach to reinstall cell cycle restriction to deregulated SSE versions, which highlights the importance of restricting SSE function to M-phase as their premature function during S-phase interferes with replication progression. As such, the observed function in M-phase matches the temporal regulation of the catalytic activity of Mus81-Mms4 and Yen1 which has been shown to be high in M-phase. For Mus81-Mms4, this upregulation of the catalytic activity is known to depend on phosphorylation by the cell cycle kinases CDK (cyclin-dependent kinase) and Cdc5 as well as on the formation of a scaffold protein complex. Here, we add a new kinase – the cell cycle kinase DDK (Dbf4-dependent kinase) – to this cell cycle regulatory network and gain insights into the interplay between the regulatory mechanisms involved. We establish that the two regulatory pathways, phosphorylation and scaffold protein complex formation, are highly interdependent and imply a switch-like activation mechanism. Taken together, our studies contribute to the understanding of the cell cycle regulation of Mus81-Mms4 and Yen1 and introduce an advanced toolbox of cell cycle tags which provides a technical source for studying cell cycle-regulated processes in general

Discovery and evolution of novel Cre-type tyrosine site-specific recombinases for advanced genome engineering

Tyrosine site-specific recombinases (Y-SSRs) are DNA editing enzymes that play a valuable role for the manipulation of genomes, due to their precision and versatility. They have been widely used in biotechnology and molecular biology for various applications, and are slowly finding their spot in gene therapy in recent years. However, the limited number of available Y-SSR systems and their often narrow target specificity have hindered the full potential of these enzymes for advanced genome engineering. In this PhD thesis, I conducted a comprehensive investigation of novel Y-SSRs and their potential for advancing genome engineering. This PhD thesis aims to address the current limitations in the genetic toolbox by identifying and characterizing novel Cre-type recombinases and demonstrating their impact on the directed evolution of designer recombinases for precise genome surgery. To achieve these aims, I developed in a collaboration a comprehensive prediction pipeline, combining a rational bioinformatical approach with knowledge of the biological functions of recombinases, to enable high success rate and high-throughput identification of novel tyrosine site-specific recombinase (Y-SSR) systems. Eight putative candidates were molecularly characterized in-depth to ensure their successful integration into future genome engineering applications. I assessed their activity in prokaryotes (E. coli) and eukaryotes (human cell lines), and determined their specificity in the sequence space of all known Cre- type target sites. The potential cytotoxicity associated with cryptic genomic recombination sites was also explored in the context of recombinase applicability. This approach allowed the identification of novel Y-SSRs with distinct target sites, enabling simultaneous use of multiple Y-SSR systems, and provided knowledge that will facilitate the assignment of novel and known recombinases to specific uses or organisms, ensuring their safe and effective implementation. The introduction of these novel Y-SSRs into the genome engineering toolbox opens up new possibilities for precise genome manipulation in various applications. The broader targetability offered by these enzymes could accelerate the development of novel gene therapies, as well as advance the understanding of gene function and regulation. Moreover, these recombinases could be used to design custom genetic circuits for synthetic biology, allowing researchers to create more complex and sophisticated cellular systems. Finally, I introduced the novel Y-SSRs into efforts aimed at developing designer recombinases for precise genome surgery, demonstrating their impact on accelerating the directed evolution process. Therapeutically relevant recombinases with altered DNA specificity have been developed for excision or inversion of specific DNA sequences. However, the potential for evolving recombinases capable of integrating large DNA cargos into naturally occurring lox-like sites in the human genome remained untapped so far. Thus, I embarked on evolving the Vika recombinase to mediate the integration of DNA cargo into a native human sequence. I discovered that Vika could integrate DNA into the voxH9 site in the human genome, and then, I enhanced the process through directed evolution. The evolved variants of Vika displayed a marked improvement in integration efficiency in bacterial systems. However, the translation of these results into mammalian systems has not yet been entirely successful. Despite this, the study laid the groundwork for future research to optimize the efficiency and applicability of Y-SSRs for genomic integration. In summary, this thesis made significant strides in the identification, characterization, and development of novel Y-SSRs for advanced genome engineering. The comprehensive prediction pipeline, combined with in-depth molecular characterization, has expanded the genetic toolbox to meet the growing demand for better genome editing tools. By exploring efficiency, cross-specificity, and potential cytotoxicity, this research lays the foundation for the safe and effective application of novel Y-SSRs in various therapeutic settings. Furthermore, by demonstrating the potential of these recombinases to improve efforts in creating designer recombinases through directed evolution, this research has opened new avenues for precise genome surgery. The successful development and implementation of these novel recombinases have the potential to revolutionize gene therapy, synthetic biology, and our understanding of gene function and regulation

Role of DDK kinase in DNA double-strand break repair and insights into the DDK-Cdc5/PLK1 kinase complex

The eukaryotic cell cycle consists of an ordered sequence of tightly regulated events to restrict specific activities within specific cycle phases. Key regulators are cell cycle kinases. Budding yeast harbor three essential cell cycle kinases conserved in humans: Dbf4-dependent kinase Cdc7 (DDK), Cyclin-dependent kinase (CDK) and the single yeast Polo-like kinase, Cdc5 (PLK1 in human). DNA double-strand breaks (DSBs) are a severe form of DNA damage. Two main pathways evolved for the repair of such toxic lesions are homologous recombination (HR) and non-homologous end joining (NHEJ). HR often uses the homologous sequence of the sister chromatid as template for error-free DSB repair. Therefore, HR is upregulated in S, G2 and M phase when a sister chromatid is present, while NHEJ is the preferred repair pathway in G1. The crucial switch from repair via NHEJ to HR is considered to be the processing of the broken ends during DNA end resection, which primes for repair by HR and inhibits repair by NHEJ. Part of this regulation comes from CDK phosphorylation of Sae2-MRX (CtIP-MRN in human), which initiates DNA end resection. However, it is increasingly clear that additional cell cycle regulated mechanism might be involved in the regulation of DNA end resection initiation. To identify novel functions of DDK, we performed phosphoproteomic experiments and discovered that DDK phosphorylates several proteins involved in DSB repair via HR, among which also factors involved in DNA end resection. We therefore followed a first line of research focused on the possible role of DDK in regulating DNA end resection. We showed that DDK is required for resection and HR, unveiling a previously unknown role of DDK in the cell cycle regulation of DSB repair. Mechanistically, we focused on phosphorylation of Sae2. We showed DDK-dependent phosphorylation of Sae2 in vivo and in vitro, and found that via phosphorylation of Sae2, DDK can stimulate the nucleolytic activity of the Sae2-MRX complex. Given the importance of DDK as regulator of resection, we tested if we could bypass the cell cycle regulation of DNA end resection and HR by forcing DDK expression in G1 cells. We observed that DDK expression in G1 cells lead to premature phosphorylation of Sae2, and by performing DNA end resection and HR assays we observed that synthetic activation of DDK in G1 leads to a moderate activation of HR, highlighting the central role of DDK in DSB repair pathway choice. Cell cycle kinases also regulate the resolution of recombination structures by the Mus81- Mms4 nuclease during late steps of HR. DDK and Cdc5 can physically interact with each other and it was previously shown that this two-kinase complex is required for phosphorylation and activation of Mus81-Mms4. In a second project we therefore focused on the DDK-Cdc5 kinase complex and how it works as two-kinase complex to phosphorylate Mus81-Mms4 and other proteins. In a candidate approach, we identified a novel phosphorylation substrate of the kinase complex, the DNA replication factor Sld2, suggesting the DDK-Cdc5 complex might be a more general regulator of M phase. We developed protocols to purify to homogeneity from yeast cells the DDK-Cdc5 complex and the single kinases. Through a series of in vitro experiments, we showed that the DDK- Cdc5 complex was overall active as well as the single kinases. Different phosphorylation substrates were specifically phosphorylated by either one of the two kinases, either when on their own or as part of the complex, suggesting that within the complex each of the kinases could act as scaffold or adaptor for substrates. Lastly, we showed that also the human proteins DDK and PLK1 (human ortholog of Cdc5) physically interact, indicating that the DDK-Cdc5/DDK-PLK1 complex is an evolutionary conserved composite cell cycle regulator. Taken together, the work presented in this thesis uncover a novel role of DDK in regulating DSB repair and offer insights into the evolutionary conserved DDK-Cdc5 kinase complex

Engineering Proteins by Domain Insertion

Protein domains are structural and functional subunits of proteins. The recombination of existing domains is a source of evolutionary innovation, as it can result in new protein features and functions. Inspired by nature, protein engineering commonly uses domain recombination in order to create artificial proteins with tailor-made properties. Customized control over protein activity, for instance, can be achieved by harnessing switchable domains and functionally linking them to effector domains. Many natural protein domains exhibit conformational changes in response to exogenous triggers. The insertion of light-switchable receptor domains into an effector protein of choice, for instance, allows the control of effector activity with light. The resulting optogenetic proteins represent powerful tools for the investigation of dynamic cellular processes with high precision in time and space. On top, optogenetic proteins enable manifold biotechnological applications and they are even considered potential candidates for future therapeutics. In this study, we first focused on CRISPR-Cas9 genome editing and applied a domain insertion strategy to genetically encoded inhibitors of the CRISPR nuclease from Neisseria meningitidis (NmeCas9), which due to its small size and high DNA sequence-specificity is of great interest for CRISPR genome editing applications. Fusing stabilizing domains to the NmeCas9 inhibitory protein AcrIIC1 allowed us to boost its inhibitory effect, thereby yielding a potent gene editing off-switch. Furthermore, the insertion of the light-responsive LOV2 domain from Avena sativa into AcrIIC3, the most potent inhibitor of NmeCas9, enabled the optogenetic control of gene editing via light-dependent NmeCas9 inhibition. Further investigation of the engineered inhibitors revealed the potential these proteins could have with respect to safe-guarding of the CRISPR technology by selectively reducing off-target editing. The laborious optimization of the engineered CRISPR inhibitors necessary by the time motivated us to more systematically investigate possibilities and constraints of protein engineering by domain insertion using an unbiased insertion approach. Previously, single protein domains were usually introduced only at a few rationally selected sites into target proteins. Here, we inserted up to five structurally and functionally unrelated domains into several different candidate effector proteins at all possible positions. The resulting libraries of protein hybrids were screened for activity by fluorescence-activated cell sorting (FACS) and subsequent next-generation sequencing (Flow-seq). Training machine learning models on the resulting, comprehensive datasets allowed us to dissect parameters that affect domain insertion tolerance and revealed that sequence conservation statistics are the most powerful predictors for domain insertion success. Finally, extending our experimental Flow-seq pipeline towards the screening of engineered, switchable effector variants yielded two potent optogenetic derivatives of the E. coli transcription factor AraC. These novel hybrids will enable the co-regulation of bacterial gene expression by light and chemicals. Taken together, our study showcases the design of functionally diverse protein switches for the control of gene editing and gene expression in mammalian cells and E. coli, respectively. In addition, the generation of a large domain insertion datasets enabled - for the first time - the unbiased investigation of domain insertion tolerance in several evolutionary unrelated proteins. Our study showcases the manifold opportunities and remaining challenges behind the engineering of proteins with new properties and functionalities by domain recombination

Visualization and manipulation of repair and regeneration in biological systems using light

Tissue repair after an injury is a fundamental process in biomedicine. It can involve regeneration, which uses new growth to restore tissue function. The interest in repair and regeneration is motivated by the desire to treat injuries and diseases and has attracted researchers for centuries. In the last decades, it evolved in the field of regenerative medicine, which has the ultimate goal of providing strategies for regenerating human cells, tissues, or even organs, for instance, via engineering principles. Already since the first experiments on regeneration by Abraham Trembley, novel findings in biomedicine, repair, and regeneration have been enabled or accompanied by research in optics, for example, on the development of novel microscopy techniques. Nowadays, novel optical techniques are advancing, which allow to understand the role of single cells in tissue repair processes. Moreover, repair processes within cells can be visualized and manipulated. Ultimately, optics can provide enabling techniques for regenerative therapies. This habilitation thesis aims to present several of these advances. On a single cell level, femtosecond laser nanosurgery was used to target specific intracellular structures during concurrent imaging in vitro. The relation of femtosecond laser nanosurgery to the cell state and cellular staining was investigated. Manipulation of single Z-discs in cardiomyocytes using a femtosecond oscillator laser system was accomplished, which allows to better elucidate the role of a single Z-disc in cardiomyocyte function. In particular, measurements on cell survival, (calcium-) homeostasis, and morphology yielded only minor deviations from control cells after single Z-disc ablation. A reduction in force generation was elucidated via traction force microscopy and gene expression level changes, for instance, an upregulation of -actinin were examined. Additionally, light-based systems to influence single cells in their alignment or to trigger single cells, for example, to activate other cells via optogenetics were applied. On the tissue scale, imaging via confocal microscopy or multiphoton microscopy has been applied for various contexts of regenerative approaches. Furthermore, a fiber-based imaging approach, which could later be used for longitudinal imaging in vivo and builds upon a fluorescence microscope system and an imaging fiber bundle in combination with reconstruction via a neural network, was developed. As another imaging strategy, an abdominal imaging window served to image the mouse liver in vivo via multiphoton microscopy in successive imaging sessions. Manipulation in tissue was applied in colonoids, which resemble the structure of the colon on an in vitro scale, and revealed different cell dynamics dependent on the location of the damage. In particular, activation of the Wnt signaling pathway after crypt damage was observed. Cell ablation via a femtosecond laser amplifier system during concurrent two-photon microscopy was also established during in vivo liver imaging to study micro-regenerative processes. Furthermore, laser-based delivery processes with novel materials or in the context of genome editing using CRISPR/Cas9 technology were investigated as enabling technologies for regenerative medicine. In conclusion, this thesis addresses the question of how optics can help to illuminate future directions in research on tissue repair and regeneration, as well as, regenerative therapies by addressing (longitudinal) imaging in a complex environment, sophisticated cell-manipulation strategies, and the application of novel materials for laser-based delivery

Engineering Controllable And Efficient Base Editors By Targeted Manipulation Of Dna Deaminases

Base editors (BEs) combine DNA deaminase mutator activity with CRISPR-Cas localization to create targeted point mutations in genomic DNA. Current approaches with BEs have enabled single-base alterations for several applications, including modeling and correction of disease alleles, crop engineering, and gene diversification. However, two major challenges limit their applicability: (1) moderate editing efficiency and (2) off-target mutagenesis of DNA and RNA. Here, we leverage our mechanistic knowledge into DNA deaminases to separately address both challenges. For one, nature has evolved DNA deaminases with suboptimal activity to achieve their role in immunity and minimize genomic instability. By deriving and characterizing hyperactive deaminases, I revealed intrinsic deaminase activity as a rate-limiting step in the base-editing reaction. Interestingly, hyperactive deaminases also had a broadened activity window, revealing a tradeoff between efficiency and precision. By harnessing their broad activity and skewing repair, we developed novel diversifying BEs that generate simultaneous C\u3eT and G\u3eA mutations efficiently over an expanded editing window of more than 65 bp. Second, DNA deaminases are highly regulated to achieve purposeful mutagenesis in physiological settings. Inspired by nature, I aimed to build regulatory control into the activity of DNA deaminases by splitting the enzyme into two inactive fragments, whose reapproximation reconstitute activity. This finding allowed me to develop small-molecule-inducible split-engineered base editors, which show decreased off-target editing when compared to intact BEs and newly enable temporal control over precise genome editing. Third, understanding the mechanistic basis for the preferential targeting of deaminases for DNA over RNA could provide means for engineering variants with decreased reactivity towards RNA. Thus, we developed biochemical assays to characterize the activity of AID/APOBEC enzymes on both substrates. Focusing on APOBEC3A, we establish the target base as a major determinant of selectivity and demonstrate that although overall deamination is greatly reduced in RNA, there is a strong selectivity for idealized substrates. Altogether, my results offer mechanistic insights into the incorporation of DNA deaminases in BEs, facilitating the development of enhanced base editing tools for diverse applications

Applications of genome editing tools in drug discovery and basic research

Since the discovery of the DNA double helix, major advances in biology have been; the development of recombinant DNA technology in the 1970s, methods to amplify DNA and gene targeting technology in the late 1980s. In organisms such as yeast and mice, the ability to accurately add or delete genetic information transformed biology, allowing an unmatched level of precision in studies of gene function. But, the ability to easily and specifically edit the genetic material of other cells and organisms remained impossible until recently for molecular biologists. The recent advent of programmable nucleases has dramatically changed the efficiency and speed of genome manipulation in several model organisms including cultured cells, as well as whole animals and plants. These tools opened up a powerful technique for biology research now called “genome editing” or “genome engineering” (Carroll, 2011; Hsu et al., 2014; Kim and Kim, 2014). In the first half of my doctoral studies, I developed genome-editing strategies to discover drug targets for a rare genetic disease called Friedreich’s Ataxia. Friedreich’s Ataxia (FRDA) is a neurodegenerative disease caused by deficiency of the mitochondrial protein frataxin (FXN) (Campuzano et al., 1997). This deficiency results from an expansion of a trinucleotide GAA repeat in the first intron of the FXN gene (Campuzano et al., 1996; Durr et al., 1996). Therapeutics that reactivate FXN gene expression are expected to be beneficial to FRDA patients (Gottesfeld, 2007). However, high-throughput screening (HTS) for FXN activators has so far met with limited success because current cellular models do not accurately assess endogenous FXN gene regulation. Here I used genome-editing technologies to generate a cellular model in which a luciferase reporter is introduced into the endogenous FXN locus. Using this system in a high-throughput genomic screen, we discovered novel inhibitors of FXN-luciferase expression. I confirmed that reducing expression of one of these inhibitors, PRKD1, led to an increase in FXN expression in FRDA patient fibroblasts (Villasenor et al., 2015). We then used reprogramming technologies to create a disease-relevant situation and test small molecules that specifically modulate PRKD1. We found that WA-21-JO19, a chemical inhibitor of PRKD1, increases FXN expression levels in iPSC-derived FRDA patient neurons. This approach, developed at the interface between academic and pharmaceutical research, demonstrates how a combination of genome editing, cellular reprogramming, and high-throughput biology can generate an effective novel drug discovery platform. In the second part of my doctoral work, we developed an interface between genome editing and proteomics to isolate native protein complexes produced from their natural genomic contexts. In many biological processes, proteins act as members of protein complexes. Understanding the molecular composition of protein complexes is a key task towards explaining their function in the cell. Conventional affinity purification followed by mass spectrometry analysis is a broadly applicable method to decipher molecular interaction networks and infer protein function. However, traditional affinity purification methods are limited by a number of factors such as antibody specificity and are sensitive to perturbations induced by overexpressed target proteins. Here, we combined genome editing with tandem affinity purification to circumvent current limitations. I uncovered subunits and interactions among well-characterized complexes and report the isolation of novel Mettl3-binding partners. The multi-protein complex composed of two active methyltransferases Mettl3 and Mettl14 mediates methylation of adenosines at position N6 on RNA molecules (Bokar et al., 1994; Bokar et al., 1997; Liu et al., 2014). N6-methyladenosine is the most abundant internal modification in eukaryotic mRNA and is often found on introns, which implies that methylation occurs co-transcriptionally (Fu et al., 2014). My work identified a set of nuclear RNA binding proteins, which specifically interact with the Mettl3-Mettl14 complex. We are currently testing the ability of these factors to function as “recruiters” of the Mettl3-Mettl14 complex to nascent mRNAs in the cell nucleus. In summary, our approach solidly establishes how a combination of genome editing and proteomics can simplify explorations of protein complexes as well as the study of post-translational modifications. In addition, this approach opens up new opportunities to study native protein complexes in a wide variety of cells and model organisms and will likely enable the systematic investigation of mammalian proteome function

Combining CRISPR-Cas9 and Proximity Labeling to Illuminate Chromatin Composition, Organization, and Regulation

A bacterial and archaeal adaptive immune system, clustered regularly interspaced short palindromic repeats and CRISPR-associated proteins (CRISPR-Cas), has recently been engineered for genome editing. This RNA-guided platform has simplified genetic manipulation and holds promise for therapeutic applications. However, off-target editing has been one of the major concerns of the commonly used Streptococcus pyogenes Cas9 (SpyCas9). Despite extensive enzyme engineering to reduce off-target editing of SpyCas9, we have turned to nature and uncovered a Cas9 ortholog from Neisseria meningitidis (Nme) with high fidelity. In the first part of my thesis, we have systematically characterized Nme1Cas9 for engineering mammalian genomes and demonstrated its high specificity by genome-wide off-targeting detection methods in vitro and in cellulo, and thus provided a new platform for accurate genome editing. Due to its flexibility, CRISPR is becoming a versatile tool not only for genome editing, but also for chromatin manipulation. These alternative applications are possible because of the programmable targeting capacity of catalytically dead Cas9 (dCas9). In the second part of my thesis, we have combined dCas9 with the engineered plant enzyme ascorbate peroxidase (APEX2) to develop a proteomic method called dCas9-APEX2 biotinylation at genomic elements by restricted spatial tagging (C-BERST). Relying on the spatially restricted, fast biotin labeling of proteins near defined genomic loci, C-BERST enables the high-throughput identification of known telomere- and centromere- associated proteomes and novel factors. Furthermore, we have extended C-BERST to map the c-fos promoter and gained new insights regarding the dynamic transcriptional regulation process. Taken together, C-BERST can advance our understanding of chromatin regulators and their roles in nuclear and chromosome biology

Guidelines for DNA recombination and repair studies: Cellular assays of DNA repair pathways

Understanding the plasticity of genomes has been greatly aided by assays for recombination, repair and mutagenesis. These assays have been developed in microbial systems that provide the advantages of genetic and molecular reporters that can readily be manipulated. Cellular assays comprise genetic, molecular, and cytological reporters. The assays are powerful tools but each comes with its particular advantages and limitations. Here the most commonly used assays are reviewed, discussed, and presented as the guidelines for future studies.European Research Council ERC2014-ADG669898 TARLOOPMinisterio de Economía y Competitividad BFU2016-75058-PJunta de Andalucía BIO123