Publications

Programmable nucleases have transformed genome editing, enabling precise DNA modification for both research and therapeutic applications. However, ensuring that these tools cut only at their intended target—and not elsewhere in the genome—remains a key challenge, particularly for clinically relevant platforms where accurate off-target detection is essential.

In our latest collaboration with the Genome Engineering and Measurement Lab (GEML) and Allogene Therapeutics, we introduce DisTAL-Seq, a method that enables genome-wide detection of TALEN-induced DNA double strand breaks directly in human cells. The experimental work was led by research technician Lena Kobel. This approach builds on the principles of DISCOVER-Seq and incorporates analysis logic tailored to TALEN binding architecture, including variable repeat specificity, cleavage offset, and dimerization behavior.

Using DisTAL-Seq, we identified and validated editing sites across different TALEN designs and primary human T-cell donors, providing a systematic view of TALEN specificity in clinically relevant contexts.

These results expand the genome-wide profiling approaches developed at GEML and provide a framework for evaluating the safety and performance of genome editing nucleases as they move towards therapeutic applications.

For more detail, check out our paper in Molecular Therapy: Nucleic Acids.

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Stimulation of the innate immune system by foreign RNA triggers a strong interferon response that helps cells defend against infection. However, this powerful defense mechanism can also lead to cell death if not properly controlled. How do cells maintain this delicate balance?

Our latest breakthrough was led by postdoc Eric Aird in collaboration with the Hale Lab (University of Zurich), Recher Lab (University of Basel, University Hospital Basel), Jackson Lab (UCAM) and University of Kuwait.

Using genome-wide CRISPR screens and follow-up cellular biochemistry experiments, we discovered that speckled protein 110 (SP110) functions as a key safeguard against cell death triggered by interferon signaling.

Mechanistically, the study revealed that SP110 interacts with the nuclear body protein SP100 to regulate the disassembly of nuclear promyelocytic leukemia (PML) bodies. Loss of SP110 made cells highly sensitive to interferon stimulation, led to mitotic retention of SP100 and PML bodies, which associated with and perturb segregating chromosomes, leading to micronucleus formation, DNA damage and genotoxic cell death. Conversely, restoring SP110 protected cells from this lethal response. The SP100-SP110 axis is molecularly achieved by newly described functions for the SP100 and SP110 CARD domains that mediate assembly and disassembly of SP100 oligomers. By controlling this process, SP110 helps maintain a balance between effective immune signaling and cell survival.

These findings highlight regulated disassembly of phase-separated biomolecular bodies as essential for cell health and that its failure may contribute to diverse human diseases.

For more details, please visit Nature Cell Biology!

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The DNA damage response (DDR) protects the genome from myriad insults. Indeed, because endogenous damage is an existential and continual threat, cells achieve robustness by engaging multiple overlapping pathways to deal with it. In the context of cancer, these relationships can present therapeutic opportunities because DDR-deficient tumors often rely on backup repair mechanisms for survival. However, addressing this complexity is a daunting challenge because gene functions in essential DNA repair processes can be masked and overlooked when pathways use completely different mechanisms to complement one another. But which DDR gene interactions are essential for cell survival during normal homeostasis? Our latest CRISPR interference (CRISPRi) screen has the answers!

In our latest paper, we performed the most systematic genetic interaction investigation of the human DDR to date. This exciting new work was driven by postdoc John Fielden and PhD student Sebastian Siegner in collaboration with the Jackson  (UCAM), Cejka (USI) and Jost  (HMS) labs.

We tested 150,000 genetic interactions, asking which DDR genes have synthetic lethal interactions with each other. We uncovered previously unknown connections between DNA repair factors as well as interactions that may be clinically exploitable. All these interactions can be browsed on our user-friendly website, SPIDRweb.

 For deep mechanistic studies, we prioritized two of the strongest novel synthetic interactions: WDR48:LIG1/FEN1 and FANCM:SMARCAL1. First, we found that WDR48 partners with USP1 to prevent PCNA degradation in cells lacking either FEN1 or LIG1, two enzymes which ordinarily prevent the accumulation of DNA nicks, gaps, and single-strand breaks. Second, we revealed that FANCM and SMARCAL1, two well-known but previously unconnected DNA translocases, have overlapping roles in unwinding DNA secondary structures that form at TA-rich repeats. In doing so, they effectively shield the genome from catastrophic fragmentation by the ERCC1-ERCC4 nuclease complex.

From a clinical perspective, our data suggest that already existing USP1 inhibitors may synergize with chemotherapies that induce DNA gaps, such as ATR and WEE1 inhibitors. Moreover, FANCM and SMARCAL1 are mutated in breast cancers and glioblastomas, respectively, pinpointing them as promising targets for drug development.

Overall, our work reveals new aspects of DDR biology and suggests multiple targets for synthetic lethality-based cancer therapy. We anticipate that our genetic interaction map will reveal further insights into fundamental DDR biology as well as cancer cell-specific vulnerabilities and candidate drug targets.

For more info check out our new paper in Nature!

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Proteins are essential building blocks of life, but they can become toxic to our cells if damaged, for example under oxidative stress. In turn, human cells selectively remove damaged proteins to maintain a healthy proteome. But how can cells identify individual damaged proteins among thousands of intact ones? An old hypothesis states that cells scan the proteome for chemical modifications that occur, for example, when proteins break. Are you interested in learning more about how cells combat chemical protein damage?

Have a look at our newest research breakthrough, led by postdoc Matthias Muhar in a collaboration with Jakob Farnung from the Bode group (D-CHAB) as well as with Jessberger group (UZH), Jinek group (UZH), Mann group (Max Plank Institute of Biochemistry) Germany   and Schulman group (Max Plank Institute of Biochemistry, Germany).

In this study, using a semi-synthetic chemical biology approach coupled to cellular assays, we found that C-terminal amide-bearing proteins (CTAPs) are rapidly cleared from human cells.

To identify the cellular machinery underlying CTAP clearance, we utilized a genome-wide CRISPR screen for genes that are responsible for specific degradation of C-terminally amidated proteins. We identified SCF–FBXO31 ubiquitin ligase as a key reader of C-terminal amides, marking CTAPs for proteasomal degradation. With a conserved binding pocket, FBXO31 exhibits remarkable selectivity, binding C-terminal peptides with amides while excluding non-modified proteins. This mechanism allows cell to remove CTAPs, which form when proteins break under under oxidative stress. Intriguingly, a human mutation linked to neurodevelopmental disorders alters FBXO31’s substrate recognition, leading to toxicity. These findings suggest CTAPs may represent a new class of modified amino acid degrons (MAADs) that mark proteins for removal by reader proteins and downstream effectors, offering insights into selective surveillance of chemically damaged proteins.

In conclusion, this research uncovered new signals for protein clearance and advanced our understanding of cellular protein quality control.

For more info check out our new paper in Nature!

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Genome editing creates double-strand breaks (DSBs) that can be repaired through either non-homologous end joining (NHEJ), microhomology-mediated end joining or homology-directed repair (HDR). While NHEJ is quick but error-prone, HDR uses a DNA template for precise edits, allowing targeted changes from single-nucleotide fixes to large gene insertions, greatly benefiting biomedical research and therapies. However, HDR is a relatively inefficient process, and ongoing efforts aim to improve its efficiency, including the use of small molecule inhibitors targeting DNA repair. One such highly potent inhibitor, AZD7648, selectively targets DNA-PKcs, redirecting DNA repair from the error-prone NHEJ pathway to the more precise HDR pathway in both transformed cell lines and primary human cells. However, the potential unintended consequences of its use in genome editing remain largely unexplored.

Are you curious to find out more?

Check out our recent advancement, led by postdoc Grégoire Cullot in a collaboration with the Gehart group (IMHS), Cathomen group (University of Freiburg, Germany) and the Gene Therapy research group of CSL Behring.

This work demonstrated that genome editing with a single Cas9-induced DSB in combination with AZD7648 leads to an increase in HDR, but this was accompanied by Cas9-induced genomic instability at on-target sites, where small-scale NHEJ outcomes were transformed into larger genetic alterations that cannot be detected by short-read sequencing.

Through the use of long-read sequencing, droplet digital PCR (ddPCR) for copy number analysis, single-cell RNA sequencing, and unbiased translocation detection, we discovered that AZD7648 significantly amplifies the frequency of kilobase-scale deletions, chromosome arm loss, and translocations across various cell types.

More broadly, genome editing-induced large-scale genomic alterations might still be largely underestimated. Indeed, these large-scale genomic alterations evade classical genome editing detection assays, typically short-read next-generation sequencing (NGS) and necessitate specific techniques that are not currently commonly used in the genome editing field. This means that clinical genome editing groups might be unaware of potential induced genomic instability and safety risks. Of note, AZD7648 is being tested by many clinical genome editing groups, however our results urge caution when deploying it during genome editing and reinforce the need to investigate genetic outcomes beyond those accessible to short-read target amplicon next-generation sequencing.

For more info check out our new paper in Nature Biotechnology!

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Cells constantly experience DNA damage throughout their lifespan due to a variety of endogenous and exogenous factors. Among the different types of DNA damage, double-stranded breaks (DSBs) are particularly severe and can be detrimental if left unrepaired. Eukaryotes have developed mechanisms to detect and repair such damage, known as the DNA damage response (DDR), which involves hundreds of key players and supporting factors. Are you curious to discover more about DDR? Have a look at latest breakthrough,  led by postdoc Erman Karasu in a collaboration with the Jonas group (IMBB) and Zavolan group (University of Basel).

By utilizing a CRISPR screening system to assay about 18,000 gene knockdowns, we identified new players influencing homology-directed repair (HDR). Among these, we focused on CCAR1, an enigmatic gene whose role in the DDR was not well understood. We found that reducing CCAR1 levels impaired both HDR and the repair of interstrand crosslinks, similar to what occurs when the Fanconi anemia (FA) pathway is inactivated. Interestingly, CCAR1 deficiency leads to FANCA protein depletion without affecting FANCA mRNA or other FA gene mRNAs levels. Instead, CCAR1 ensures FANCA mRNA splicing by suppressing a poison exon inclusion. CCAR1 binds to U2-type spliceosome and acts on multiple splicing sites. This demonstrates that CCAR1 is crucial for proper mRNA splicing. Further analysis showed that CCAR1’s role in mRNA splicing extends beyond FANCA, affecting many genes and ensuring that their mRNAs are correctly processed in both mouse and human cells. Our work highlights CCAR1’s unexpected role in maintaining accurate mRNA splicing, which is essential for proper protein function.

For more info check out our new paper in Molecular Cell!

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Erythrocytes, or red blood cells, carry hemoglobin and circulate throughout the body to supply oxygen. β-hemoglobinopathies, such as sickle cell disease and β-thalassemia, are the most common genetic diseases worldwide and are caused by mutations affecting the structure or production of β-globin subunits in adult hemoglobin. These conditions result in anemia and organ damage, and available treatment options are limited. Stem cell transplantation is currently the only curative approach, although its feasibility relies on the availability of a suitable donor.

Hemoglobin is a tetrameric protein composed of 2 α-like (HBA) and 2 β-like subunits (HBB). Hemoglobin A1 (HbA1) constitutes 97% of adult hemoglobin, while Hemoglobin A2 (HbA2) makes up 2-3%. HbA2 is composed of two α-globin subunits and two δ-globin (HBD) subunits. HBD is a homologous to HBB gene, but with much lower expression compared to HBB due to a weak promoter.  Currently, many efforts are focused on increasing fetal hemoglobin (HbF) to treat the β-hemoglobinopathies. But HbA2 is more similar to HbA1 and is already expressed at low levels in all adult red blood cells. What if we were to increase HbA2 levels? Could they potentially compensate for beta-globin deficiency? Can genome editing technologies be used to boost transcriptional activity of the endogenous HBD promoter to increase HbA2 levels? Mandy Boontanrart, a Postdoc in our lab, was eager to discover the answers to these questions.

Using CRISPR-Cas9 genome editing, we inserted various transcription factor binding sequences into the endogenous HBD promoter. Team efforts yielded positive results as we successfully increased the transcriptional activity in HUDEP-2 immortalized erythroid progenitor cells, resulting in a significant upregulation of HBD expression. Despite roughly equal homology-directed repair rates between all promoter designs, we observed a significant increase in HBD only for the design with all three elements (KLF1, β-DRF, and TFIIB). We next explored whether endogenous editing of the HBD promoter can be accomplished in bone marrow stem cells. We found up to 46% HBD expression in clonal populations. We also tested a small molecule drug that enhances HDR outcomes by inhibiting the NHEJ pathway and observed an increase in the percent of HDR alleles in pooled edited bone marrow stem cells.

While our findings provide key mechanistic insight into the globin gene regulation, several questions remain to be tackled.  Is heterozygous knock-in of the promoter design in β-hemoglobinopathy cells is sufficient to ameliorate disease phenotypes? What is the safety profile of this strategy?

Overall, our work is a promising approach for restoring hemoglobin levels in red blood cells. This strategy might open new therapeutic avenues for to treating beta-hemoglobinopathies in the future.

For more, check out our paper, it is now out in Elife!

Note: Excitingly, Mandy is now leading an ETH spin-off, building upon the findings of the paper, check out their brand-new website https://www.ariyabio.ch/!

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Cas-mediated genome editing technology holds great promise as a curative treatment for a number of genetic diseases. Conventional CRISPR-Cas genome editing induces DNA damage (double stranded DNA breaks) and relies on the cellular DNA repair system to yield the desired repair outcome. This works quite well in cells with a fully operational DNA repair machinery. However, in Fanconi Anemia (FA), a genetic disorder associated with bone marrow failure and cancer predisposition, DNA repair is defective due to the gene mutations causing the disorder. Homology directed repair (HDR)-based editing strategy as an option to correct FA mutations is pretty inefficient (Richardson et al. Nature Genetics 2018). But is it possible to avoid DNA cleavage, and instead use recently developed genome editing systems such as base editing? Can base editing reverse the effects of FA mutations? New work led by a postdoc Erman Karasu (co-corresponding author) and PhD student Sebastian Siegner together with Alexandra Clemens at ETHZ and Laura Ugalde from Paula Rio’s lab shows that this is possible, even in bone marrow stem cells from FA patients.

In this proof-of-concept study, we find that base editing can indeed restore the function of FA bone marrow stem cells. First, the team went through cycles of optimization for the conditions (base editor construct, vector type, guide RNA format, delivery) in cell lines from multiple FA patients. The developed approach effectively corrected FA mutations in both patient-derived cell lines and bone marrow stem cells from FA patients, leading to restored FANCA expression and functional FA pathway and phenotypic resistance to crosslinking agents.

An obvious question that comes up: how safe is this editing approach? To answer this question, the team predicted possible off-targets and measured the editing outcomes in > 60 sites across multiple base editors. Unintended modifications were detected at a single site with one guide RNA, but a guide RNA targeting the most prevalent FA mutation had no detected off-targets.  Nevertheless, un-biased off- target identification using genome or RNA-sequencing will be the next step in the preclinical validation of base editing as an approach to cure FA.

Altogether, this work highlights base editors as a feasible editing strategy in FA and brings us one step closer to the future clinical implementation of base editing not only in FA, but also in other genetic diseases.

Check out our paper, now out in Nature Communications!

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Are you interested in autophagy of really big things in the cells, such as organelles?  Are you wondering how cutting-edge genetic tools can accelerate your autophagy research? Jin Rui Liang (Amos), a former postdoc, now running his own group at the University of Dundee, wrote a comprehensive review on CRISPR’s impact on autophagy research. Amos outlines the major considerations for CRISPR-based genetic manipulations in autophagy, with a focus on genome-wide screening, as well as various reporter systems for high-throughput autophagy quantification. The article thoroughly summarizes all the relevant info on recently performed autophagy-related screens and their discoveries. Have a CRISPR-view on autophagy, now out in Trends in Cell Biology!

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The Coronavirus lockdowns this spring disrupted many projects and students. But when life gives you lemons, make lemonade. In our lab, almost everyone took the stay-at-home orders as motivation to learn some coding. Sebastian Siegner, a Masters’ student at the time and now recently joined for a full Ph.D., had been working on proof-of-concept experiments for therapeutic base editing and prime editing. But he was frustrated with designing base editor gRNAs and newly-described prime editor pegRNAs by hand. So he used the lockdown to write PnB Designer, which is a fast and scalable helper to design your prime and base editing experiments. Check out the paper in BMC Bioinformatics.

PnB designer can be used to design base editing gRNAs and prime editing pegRNAs in both single-edit and batch mode. You can design against arbitrary DNA sequences (copy/paste your sequence as input), or you can enter genomic coordinates of your favorite gene in your favorite genome. Several species are currently supported, from human to plant. For base editors, just choose which nucleotide change you want to make and the software will take into account both the mutation and editing window to figure out the best editor to use. For prime editing, you can test all kinds of reverse transcriptase template (RTT) and primer binding site (PBS) lengths with a click of a button. The output is a table of possible gRNAs/pegRNAs, ordered by a heuristic score.

Sebastian tested PnB Designer by designing pegRNAs to model most of the human disease-associated mutations in ClinVar using prime editing. He even varied RTT and PBS length for each of these 96,000+ mutations, figuring out good parameters to keep pegRNAs at a reasonable length while still modelling ~80% of all variants.

This was a challenging but exciting side-project for Sebastian’s Masters’ degree. He wrote PnB Designer independently during the lockdown, and the rest of the lab acted as beta testers to provide suggestions. Congrats to Sebastian on your first paper, which is already being used by several people in the Zurich area. PnB Designer is completely free to use and is hosted by the Functional Genomics Center Zurich.

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