Publications

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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When using CRISPR genome editing in stem cells, it’s far easier to break a gene with indels than to fix it with HDR. This manifests in an interesting way. If you monitor a “CD34+” population of hematopoietic stem and progenitor cells (HSPCs) from the bone marrow, indels start high and stay high but HDR alleles are lost over time. Why do these different genetic outcomes differ over time? Is HDR bad for the long-term stem cells? Or is editing in the CD34+ population actually heterogeneous, and different cells get different alleles? New work from postdoc Jenny Shin in the lab, out in Cell reports, both answers this question and finds a way to fix the problem.

Jenny and collaborators used a powerful combination of immunophenotyping, next generation sequencing, and single-cell RNA-sequencing to investigate and reprogram genome editing outcomes in subpopulations of adult human CD34+ HSPCs. These HSPCs are actually several different types of cells, including more differentiated progenitors that cycle and very “stemmy” long-term HSCs that are quiescent. The team found that there is a dramatic tension between HDR and quiescence in LT-HSCs.  Quiescent stem-enriched cells utilize NHEJ and exhibit almost no HDR. By contrast, non-quiescent cells with the same immunophenotype utilize both NHEJ and HDR. Quiescence is critical for engraftment and stem cell maintenance, so it was now clear that all cells in the CD34+ population get indels and the cycling progenitors were getting HDR alleles, but the quiescent LT-HSCs weren’t doing HDR.

Jenny then had a very creative idea. She asked if a previously reported small molecule cocktail, “XRC”, that maintains quiescence could be used after the fact to re-quiesce LT-HSCs. Using this new strategy and good timing, she found a way to get LT-HSCs with high levels of HDR by briefly allowing them to cycle during editing, and then inducing quiescence later on. This yielded a 6-fold increase in the HDR/NHEJ ratio in quiescent stem cells ex vivo and during long-term engraftment in mouse experiments. The re-quiescence strategy might in future be combined with engineered Cas9-geminin constructs that reduce NHEJ, further tipping the balance towards HDR. Jenny’s results highlight the tradeoffs between editing and fundamental cellular physiology and suggests strategies to manipulate quiescent cells for research and therapeutic genome editing. 

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When my lab published editing of the sickle beta globin gene, together with Mark Walters and David Martin, we noticed something quite unexpected. Every time we targeted beta globin, we saw fetal globin go up! We also noticed this in beta globin editing papers from the labs of Paula Cannon and Matt Porteus. Transient stress can cause blood cells to express fetal globin, so at first we thought that we were just seeing something short-lived. But it turned out that fetal globin persisted after long-term transplantation of edited human cells into a mouse model. What could be happening in these cells? Could this be a route to treat globin-related diseases such as sickle cell disease? If so, why is beta thalassemia (lack of beta globin) not automatically compensated by fetal globin expression? A new paper from PhD student Mandy Boontanrart, out now in Cell Reports, finally figures out what’s going on. This work was completely driven by Mandy, using a wide range of techniques to dig into a difficult problem.

Editing itself can be stressful to cells, but Mandy’s first big findings were that the fetal globin response is not related to the the process of editing. Doing a scan of CRISPR cutting guides through beta globin, she found that any guide targeting an exon causes fetal globin to go up. But equally effective guides that target introns have no effect. She also found that CRISPRi transcriptional repression to knockdown beta globin does increase fetal globin. Therefore, it was something special about lacking beta globin, which is known as B0-stress.

Using clones of blood precursors edited to lack beta globin, Mandy did RNA-seq during differentiation to figure out how cells responded over time. The number one pathway by far was ATF4 signalling. But in a very strange way! It turned out that ATF4 signalling actually went down in cells with beta globin knockout. This was surprising because ATF4 responds to unfolded proteins, it had been suggested that lack of beta globin would lead to alpha chain aggregation that should turn on the unfolded protein response. We were seeing the exact opposite!

This actually makes sense in the context of recent work from the lab of Gerd Blobel, who found out that knockout of the heme-responsive kinase HRI can also increase fetal globin. HRI is upstream of eIF2a and ATF4, so it seemed that free heme (caused by lack of complete adult hemoglobin tetramers) had a stronger role than the free alpha chains in our system.

But what was ATF4 doing to increase fetal globin? Mandy investigated this using a large number of isogenically-controlled ChIP-seq experiments. She found that ATF4 doesn’t bind anywhere in the globin locus, nor does it bind anywhere near BCL11A, which is one of the latest hot globin regulators. In fact, Mandy found that ATF4’s effect on fetal globin still happened in K562 cells, which don’t express any BCL11A at all. Instead, Mandy found strong evidence that ATF4 regulates Myb through binding to a known enhancer region. Myb is itself a regulator of Bcl11A, but can also regulate fetal globin through other factors such as KLF1.

As always there’s a lot more to figure out. And it’s still unclear if we can use these findings to make a difference for patients with globinopathies. But thanks to Mandy’s work, we finally know why fetal globin increases when beta globin is reduced. We also have some clues as to why beta thalassemia is not always compensated by increased fetal globin. In fact, to some extent, it is! Many people with beta thalassemia exhibit increased fetal globin, but not enough to alleviate the symptoms of complete lack of beta globin. ATF4 does many jobs, so it could be that reducing ATF4 enough to yield large amounts of fetal globin might be bad for other biologies. This also adds a note of caution to targeting HRI as a therapeutic angle to increase fetal globin, since doing so could have pleiotropic effects. But all of that remains to be tested!

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Cas9 is a great DNA cutting enzyme, but it’s also a little weird. Unlike other nucleases (such as restriction enzymes), S, pyogenes Cas9 sticks on DNA for a looooong time. In fact, it spend the same amount of time on DNA whether it is active or inactive! In a test tube, it takes hours and hours to let go of even a cleaved DNA strand. So how does genome editing even work? Does a human cell care that Cas9 it stuck on its genome? Does it have ways to knock it off?

Superstar grad student Alan Wang’s new paper, out today in Molecular Cell, solves this mystery. Previous work suggested that RNA polymerase II was capable of displacing Cas9 in vitro. But Cas9-based technologies work even when targeted to a non-transcribed region. Alan’s first hint that something interesting was going on came from work in Xenopus egg extract, in collaboration with the lab of Johannes Walter. This “frog juice” is very often used to study DNA repair and can be elegantly deconstructed to figure out biological function. Cas9 in buffer sticks on DNA for a long time, but in Xenopus extract it comes off almost immediately!

Alan took an unbiased approach to figure out what was removing Cas9 from DNA. He fused recombinant Cas9 with a promiscuous biotin ligase, bound purified Cas9-ligase  to a plasmid, and used mass spectrometry to figure out what pushed the Cas9 off the plasmid. We were pleasantly surprised to find that both subunits of a dimeric histone chaperone called FACT were top mass spec hits! Follow-up experiments showed that FACT was necessary and sufficient for displacing Cas9 from DNA substrates. FACT was responsible for turning Cas9 from a multi-turnover “classic” nuclease enzyme into a single-turnover sticky enzyme!

In human cells, FACT had several interesting effects on genome manipulation. Knocking down FACT delayed homology directed repair and altered the balance of repair outcomes. FACT knockdown increased epigenetic marking from both CRISPRi and CRISPRa constructs, and increased CRISPRi phenotypes. We attribute this to increasing the residence time at a target site: giving the effect fused to Cas9 more time to have an effect on the genome. 

The take-home is that cells are not passive players, and play a leading role in genome manipulation. The cell is just as important as the enzyme! Alan’s work starts to reveal how cells monitor their genomes during Cas9 interventions.

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When doing genome editing, fixing sequences by HDR is better than breaking them by making indels. If you really want to break something, you could even use HDR to insert a precise indel or a stop codon. Unfortunately, HDR is relatively inefficient in human cells. Single stranded oligo donors help, but editing the same locus with a double stranded plasmid DNA donor is almost always painful. What’s the bottleneck? We tried to answer this question in a new paper just out in Nature Communications.

To answer this question, Chris Richardson and Beeke Wienert led a superstar team to perform CRISPRi screening while simultaneously editing with a double stranded DNA plasmid donor. The screen itself was performed by Sharon Feng, a superstar undergraduate. Putting everything together was an exciting collaboration with the labs of Bruce Conklin and Alex Marson.

The first set of hits are known homologous recombination factors, such as BRCA1. This gives high confidence that the screen worked as expected. Surprisingly, the same Fanconi Anemia complexes that are required for single stranded oligo HDR are required for plasmid HDR. The FA pathway is thus a core regulator of all forms of HDR!

But we really wanted to know genes could increase HDR if they were removed. Knocking down a gene is hard to do in many contexts. So we focused on genes with known inhibitors. It turns out that small molecule inhibitors of CDC7 give very nice boosts in HDR from both single stranded oligo and plasmid DNA donors. This works for small changes (SNPs), medium changes (adding epitope tags) and even large cargoes (site-targeted transgenes). It also works in a variety of cell types, including hematopoietic stem cells and T cells. Not every cell is created equally, so check out the paper for detailed guidelines.

Our favorite CDC7 inhibitor is XL413, which is non-toxic and quite reversible. This distinguishes it from some other HDR-improving compounds that lead big genomic messes, including polyploidy. Delving into mechanism, CDC7 inhibition leads to loss of MCM2 phosphorylation. Because MCM2 phosphorylation is required for S phase progression, XL413 leads to a longer S phase. This is a magical phase of the cell cycle for HDR, and so our model is that XL413 increases HDR by increasing the amount of time cells are able to do HDR. We tested this with a timing experiment. Hitting cells with Cas9 and then immediately putting them in XL413 leads to increased HDR, because the cells are piling up in S phase at the same time they are repairing the Cas9 damage. But putting cells in XL413 first and then taking them out during editing leads to decreased HDR. This is because the cells all pile up S phase before editing, and then exit into HDR non-permissive cell cycle while Cas9 is doing its thing.

We hope other labs find XL413 to be useful to increase HDR. It’s not a magic bullet and seems to work especially well in hematopoietic lineages and iPSCs. If you try it out in your favorite cells, please let us know your experience!

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Did you know that cells eat their own organelles? This is best known when damaged mitochondria are degraded by autophagy (aka mitophagy). Failure to perform mitophagy can lead to diseases such as Parkinson’s. But many other organelles are also degraded by autophagy. We have been studying autophagy of the endoplasmic reticulum (ER-phagy), which is much less understood than mitophagy. Engulfing a mitochondria in an autophagosome sounds pretty straightforward, but ends up being complicated. Now imagine needing to do that for one part of the ER network! A handful of direct ER-phagy receptors are known. But these receptors are always on the ER, so it was not clear what really initiates and controls ER-phagy. How does the cell know what part to engulf? What are the signals that turn this on and off? What happens when it goes wrong?

Superstars Amos Liang (postdoc) and Emily Lingeman (PhD student) tackled this question in a big way. Using a highly sensitive fluorescent reporter for ER-phagy, they used CRISPRi to ask what genes regulate ER-phagy. The first surprise was that intact mitochondrial oxidative phosphorylation is required to successfully initiate ER-phagy. This is odd because preventing oxidative phosphorylation actually initiates bulk autophagy. But the opposite is true for ER-phagy! The second big surprise was that a weird post-translational modification called UFMylation is required for ER-phagy. Lots of mechanistic work showed how UFMylation machinery is brought to the ER surface and what gets UFMylated during ER-phagy. There are some very interesting parallels to mitophagy, but using totally different machinery. Third, many of the genes involved in ER-phagy are involved in peripheral neuropathy in humans. Since their role in ER-phagy wasn’t previously known, it wasn’t understood how they were connected to cause human disease. This work suggests that failure to do ER-phagy links them all and leads to neurodegeneration. There’s a lot going on here, so read the paper to find out more.  Congrats to Amos and Emily!

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