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Dr. Erik Willems was trained as a stem cell biologist in Brussels, Belgium where he obtained his PhD in 2007, after which he soon relocated to San Diego, California to develop his expertise in the use of pluripotent stem cells in high throughput screening assays for understanding the basic biology and disease of the developing heart at the Sanford Burnham Prebys Medical Discovery Institute. Dr. Willems then pursued his passion for the development of biotechnology tools and applications and joined Thermo Fisher Scientific in Carlsbad, California where he - as a Manager in the Cell Biology Business - currently leads pluripotent stem cell-based customer driven projects and product applications, including characterization, reprogramming, genome editing, differentiation and disease modeling with a focus on drug discovery applications. Now in the stem cell field for over 15 years, Dr. Willems published numerous peer reviewed articles including in high impact journals such as Cell Stem Cell. His key expertise encompasses pluripotent stem cell biology, differentiation, genome editing, high throughput screening and drug discovery.
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Embryonic development is a crucial period in the life of a multicellular organism, during which limited sets of embryonic progenitors produce all cells in the adult body. Determining which fate these progenitors acquire in adult tissues requires the simultaneous measurement of clonal history and cell identity at single-cell resolution, which has been a major challenge. Clonal history has traditionally been investigated by microscopically tracking cells during development, monitoring the heritable expression of genetically encoded fluorescent proteins and, more recently, using next-generation sequencing technologies that exploit somatic mutations, microsatellite instability, transposon tagging, viral barcoding, CRISPR-Cas9 genome editing and Cre-loxP recombination. Single-cell transcriptomics provides a powerful platform for unbiased cell-type classification. Here we present ScarTrace, a single-cell sequencing strategy that enables the simultaneous quantification of clonal history and cell type for thousands of cells obtained from different organs of the adult zebrafish. Using ScarTrace, we show that a small set of multipotent embryonic progenitors generate all haematopoietic cells in the kidney marrow, and that many progenitors produce specific cell types in the eyes and brain. In addition, we study when embryonic progenitors commit to the left or right eye. ScarTrace reveals that epidermal and mesenchymal cells in the caudal fin arise from the same progenitors, and that osteoblast-restricted precursors can produce mesenchymal cells during regeneration. Furthermore, we identify resident immune cells in the fin with a distinct clonal origin from other blood cell types. We envision that similar approaches will have major applications in other experimental systems, in which the matching of embryonic clonal origin to adult cell type will ultimately allow reconstruction of how the adult body is built from a single cell.
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Fragile X syndrome (FXS), the most common genetic form of intellectual disability in males, is caused by silencing of the FMR1 geneassociated with hypermethylation of the CGG expansion mutation in the 5' UTR of FMR1 in FXS patients. Here, we applied recently developed DNA methylation editing tools to reverse this hypermethylation event. Targeted demethylation of the CGG expansion by dCas9-Tet1/single guide RNA (sgRNA) switched the heterochromatin status of the upstream FMR1 promoter to an active chromatin state, restoring a persistent expression of FMR1 in FXS iPSCs. Neurons derived from methylation-edited FXS iPSCs rescued the electrophysiological abnormalities and restored a wild-type phenotype upon the mutant neurons. FMR1 expression in edited neurons was maintained in vivo after engrafting into the mouse brain. Finally, demethylation of the CGG repeats in post-mitotic FXS neurons also reactivated FMR1. Our data establish that demethylation of the CGG expansion is sufficient for FMR1 reactivation, suggesting potential therapeutic strategies for FXS.
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Rapid detection of nucleic acids is integral for clinical diagnostics and biotechnological applications. We recently developed a platform termed SHERLOCK (Specific High Sensitivity Enzymatic Reporter UnLOCKing) that combines isothermal pre-amplification with Cas13 to detect single molecules of RNA or DNA. Through characterization of CRISPR enzymology and application development, we report here four advances integrated into SHERLOCKv2: 1) 4-channel single reaction multiplexing using orthogonal CRISPR enzymes; 2) quantitative measurement of input down to 2 aM; 3) 3.5-fold increase in signal sensitivity by combining Cas13 with Csm6, an auxilary CRISPR-associated enzyme; and 4) lateral flow read-out. SHERLOCKv2 can detect Dengue or Zika virus ssRNA as well as mutations in patient liquid biopsy samples via lateral flow, highlighting its potential as a multiplexable, portable, rapid, and quantitative detection platform of nucleic acids.
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Current genome-editing systems generally rely on inducing DNA double-strand breaks (DSBs). This may limit their utility in clinical therapies, as unwanted mutations caused by DSBs can have deleterious effects. CRISPR/Cas9 system has recently been repurposed to enable targetgene activation, allowing regulation of endogenous gene expression without creating DSBs. However, in vivo implementation of this gain-of-function system has proven difficult. Here, we report a robust system for in vivo activation of endogenous target genes through trans-epigenetic remodeling. The system relies on recruitment of Cas9 and transcriptional activation complexes to target loci by modified single guide RNAs. As proof-of-concept, we used this technology to treat mouse models of diabetes, muscular dystrophy, and acute kidney disease. Results demonstrate that CRISPR/Cas9-mediated target gene activation can be achieved in vivo, leading to measurable phenotypes and amelioration of disease symptoms. This establishes new avenues for developing targeted epigenetic therapies against human diseases. VIDEO ABSTRACT.
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The CRISPR-Cas9 system has proven to be a powerful tool for genome editing allowing for the precise modification of specific DNA sequences within a cell. Many efforts are currently underway to use the CRISPR-Cas9 system for the therapeutic correction of human genetic diseases. The most widely used homologs of the Cas9 protein are derived from the bacteria Staphylococcus aureus (S. aureus) and Streptococcus pyogenes (S. pyogenes). Based on the fact that these two bacterial species cause infections in the human population at high frequencies, we looked for the presence of pre-existing adaptive immune responses to their respective Cas9 homologs, SaCas9 (S. aureus homolog of Cas9) and SpCas9 (S. pyogenes homolog of Cas9). To determine the presence of anti-Cas9 antibodies, we probed for the two homologs using human serum and were able to detect antibodies against both, with 79% of donors staining against SaCas9 and 65% of donors staining against SpCas9. Upon investigating the presence of antigen-specific T-cells against the two homologs in human peripheral blood, we found anti-SaCas9 T-cells in 46% of donors. Upon isolating, expanding, and conducting antigen re-stimulation experiments on several of these donors anti-SaCas9 T-cells, we observed a SaCas9-specific response confirming that these T-cells were antigen-specific. We were unable to detect antigen-specific T-cells against SpCas9, although the sensitivity of the assay precludes us from concluding that such T-cells do not exist. Together, this data demonstrates that there are pre-existing humoral and cell-mediated adaptive immune responses to Cas9 in humans, a factor which must be taken into account as the CRISPR-Cas9 system moves forward into clinical trials.
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Despite their fundamental biological and clinical importance, the molecular mechanisms that regulate the first cell fate decisions in the humanembryo are not well understood. Here we use CRISPR-Cas9-mediated genome editing to investigate the function of the pluripotency transcription factor OCT4 during human embryogenesis. We identified an efficient OCT4-targeting guide RNA using an inducible humanembryonic stem cell-based system and microinjection of mouse zygotes. Using these refined methods, we efficiently and specifically targeted the gene encoding OCT4 (POU5F1) in diploid human zygotes and found that blastocyst development was compromised. Transcriptomics analysis revealed that, in POU5F1-null cells, gene expression was downregulated not only for extra-embryonic trophectoderm genes, such as CDX2, but also for regulators of the pluripotent epiblast, including NANOG. By contrast, Pou5f1-null mouse embryos maintained the expression of orthologous genes, and blastocyst development was established, but maintenance was compromised. We conclude that CRISPR-Cas9-mediated genome editing is a powerful method for investigating gene function in the context of human development.
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