CRISPR

Defining Microbial and Host Pathways Driving Asymptomatic C. difficile Colonization Associated with Aging and High-Sugar Diets

SUMMARY Clostridioides difficile infection (CDI) is a leading cause of healthcare-associated diarrhea, with rising incidence in community settings and a growing burden of asymptomatic colonization. Asymptomatic car- riers, particularly among the elderly and individuals consuming high-sugar diets, represent a critical but underexplored reservoir for transmission and disease progression. This proposal introduces novel, anti- biotic-independent mouse models demonstrating that both dietary sugar and aging independently pro- mote asymptomatic C. difficile colonization. We hypothesize that these factors disrupt colonization re- sistance (CR) through distinct but overlapping microbial, metabolic, and immune pathways. In Aim 1, we will define how traditional and emerging dietary sugars alter the gut environment to permit C. difficile colonization using in vitro bioreactors and in vivo models. Aim 2 will identify age-associated changes in microbiota and mucosal immunity that impair CR, using longitudinal studies and fecal micro- biota transfer. Aim 3 will functionally validate C. difficile genes upregulated during asymptomatic carriage using CRISPR-Cas9 mutants in both sugar- and age-induced models. This integrative, multi-omics approach will uncover the mechanisms enabling asymptomatic colonization and identify microbial and host targets for intervention. The findings will inform microbiome-based strat- egies to prevent CDI in vulnerable populations and shift current paradigms in CDI risk assessment and prevention.

Mechanisms of antigen-specific T cell activation in MOGAD

PROJECT SUMMARY / ABSTRACT The overarching goal of this application is to train Dr. Carson E. Moseley, MD, PhD, who is a clinical neurologist and a research immunologist, to become an independent investigator studying and treating neuroimmunologic disorders. Myelin oligodendrocyte glycoprotein (MOG) antibody-associated disease (MOGAD) is a recently described, severe, neuroinflammatory syndrome of the central nervous system (CNS) with no approved therapies. Although MOG-specific antibodies helped define the disease, MOG antibodies alone are not clearly pathogenic and our understanding of MOGAD immunopathology is limited. CD4+ T cells are a dominant lymphocyte population in MOGAD lesions, yet the targets of T cell responses to MOG and how T and B cells interact to drive pathogenic immune response in MOGAD are unknown. This proposal uses a complementary approach of human and mouse immunology along with new technologies in T cell repertoire mapping and genome editing to dissect MOG-specific CD4+ T cell responses in MOGAD. Additionally, it will use new models to investigate how B cells promote pathogenic T cell differentiation and select pathogenic T cell receptors. The proposed training plan involves mentored training, seminars, formal learning, and advising to ensure completion of the proposed research and Dr. Moseley’s career development. He will train at UCSF, which is an outstanding institute for research and environment for physician-scientists. He will receive training in human immunology and CRISPR-based gene editing technologies. He will be mentored by Dr. Scott Zamvil, a leader in identifying antigen-specific T cell responses in neuroimmunologic disorders, and co-mentored by Dr. Alexander Marson, an expert in CRISPR gene editing to understand lymphocyte function. This application will provide Dr. Moseley with the long-term skills needed to become an independent investigator leading efforts to study and treat neuroimmunologic disorders.

Targeting disulfidptosis in cancer: mechanisms and preclinical translation

Project Summary Studying regulated cell death is critical for our understanding of cellular homeostasis and tumor suppression. We recently discovered disulfidptosis as a new form of regulated cell death induced by disulfide stress under NADPH-depleting conditions in SLC7A11-high cancer cells. However, in contrast to our deep understanding of other cell death modalities such as apoptosis and ferroptosis, the molecular and metabolic underpinnings of disulfidptosis, along with its therapeutic implications, remain largely unexplored. The objectives of this application are to elucidate the mechanisms underlying disulfidptosis and to therapeutically target this form of cell death in SLC7A11-high cancers. The proposed studies will make extensive use of human cancer cell lines and integrated human cellbased molecular analyses, including metabolomics, proteomics, CRISPR screening, and biochemical studies, to define the metabolic and signaling mechanisms governing disulfidptosis. In addition, select in vivo studies are incorporated in the therapeutic validation components of the project, where tumor growth response, systemic drug exposure and tolerability, tumor microenvironmental influences, and host immune/stromal interactions must be evaluated in an organismal context to ensure translational rigor. Alternative in vitro systems such as organoids may provide useful complementary information on tumor-intrinsic responses, but they cannot fully recapitulate the systemic metabolic stress, pharmacologic exposure, and organism-level therapeutic efficacy required for these studies. It is expected that our proposed studies will reveal novel mechanisms underlying disulfidptosis and identify effective therapies to induce this form of cell death in SLC7A11-high cancers. Our proposal is highly innovative because it focuses on a previously unexplored cell death pathway in cancer therapy. Our proposed studies will have significant impact on both our understanding of the fundamental mechanisms of disulfidptosis and our ability to target this cell death pathway in cancer treatment.

Causal mechanisms driving germline predisposition to myeloproliferative disorders

SUMMARY/ABSTRACT Although human genetic studies have indicated a significant hereditary predisposition to myeloproliferative neoplasms (MPNs) the underlying mechanisms driving the genetic risk remains unknown. Our large genome wide association study (GWAS) on MPNs identified several non-coding genetic risk loci associated with disease and implicated modulation of hematopoietic stem cell (HSC) self-renewal by the genetic variants. The long-term goal is to utilize our GWAS results to better understand MPN disease initiation and progression and draw out key unknown MPN predisposition genes. The overall objectives in this application are to elucidate the mechanisms by which MPN risk variants promote disease initiation and progression. The central hypothesis is that common genetic variants increase MPN risk by affecting regulatory elements that influence clonal expansion of HSCs carrying MPN driver mutations. The rationale for this project is that the HSC clones with most prevalent driver mutation found in MPN, JAK2V617F show individual specific growth rates and can develop into MPN or remain as clonal hematopoiesis without any consequences indicating that germline genetic factors influence this process. The central hypothesis will be tested by pursuing two specific aims: 1) To determine the mechanisms by which genetic variation at the GFI1B locus influences MPN predisposition in vivo. 2) To define upstream transcriptional mechanisms disrupted by common genetic variants that predispose to MPN. Under the first aim, a newly generated mouse model will be used to evaluate clonal expansion of JAK2V617F HSCs in the context of a germline Gfi1b enhancer deletion by in vivo competitive transplantation assays. The murine studies will be complemented by an assessment of Gfi1b allele specific clonal expansion in primary human hematopoietic stem and progenitor cells (HSPCs) engineered to carry JAK2V617F mutation. Mechanistically activated mitochondrial respiration will be examined in germline enhancer inactivated JAK2V617F HSPCs in murine models and human patient samples. For the second aim, perturbation of RUNX1 bound cis-regulatory elements by MPN risk variants will be evaluated as a mechanism of clonal expansion in MPN by using lentiviral reporter assays and endogenous CRISPR/Cas9 editing approaches in primary human HSPCs and degron tagged RUNX1 cell lines. A Runx1 haploinsufficiency mouse model will be used to assess global influences of RUNX1 transcriptional network on MPN initiation. Collectively, our proposed studies aim to bridge the gap between inherited genetic variations and the clonal expansion dynamics of MPN stem cells, shedding light on crucial factors influencing disease development. The mouse models proposed in this study provide the in vivo physiological context and functional readouts required to investigate HSC clonal expansion and MPN pathogenesis.

Linking Single-Cell Transcriptomic, Morphological, and Temporal Signatures of Vulnerability in Neurodegeneration

Neurodegeneration involves complex cellular phenotypes and molecular changes that vary widely among the cells of the nervous system. Current methodologies permit either detailed molecular profiling (e.g., single-cell transcriptomics) or functional phenotyping (e.g., live imaging of neuronal activity), but not both in the same cells. Thus, it is difficult to directly link a neuron's functional state or fate with its gene expression profile. To address this limitation, we developed an innovative technology, VISTA-FISH (Video Imaging with Spatial- Temporal Analysis by FISH), that couples prospective live-cell imaging with high-resolution spatial transcriptomic profiling of the same cells. This approach enables in situ comparisons of gene expression in neurons that exhibit divergent behaviors or outcomes. Using VISTA-FISH, we will profile iPS-derived human neurons to link single-cell gene expression, morphology, and temporal phenotypes to study molecular pathways driving resilience as well as susceptibility. After exposing neurons carrying TDP43 and C9orf72 mutations to a stimulus inducing TDP43 aggregation, we will jointly record TDP43 localization and neuron activity using live-cell microscopy, then measure single-cell gene expression of the same cells (Aim 1). We will also combine live-cell measurements of TDP43 half-life with CRISPR screening and single-cell gene expression (Aim 2). These rich datasets will enable us to determine transcriptomic changes associated with differences in protein aggregation, protein synthesis, and protein degradation in individual cells, providing an unprecedented molecular perspective on factors responsible for vulnerability and resilience to neurodegeneration.

ATPase Chromatin Remodeling Complexes as Modulators of HIV-1 Latency and Therapeutic Targets

Abstract Significance: HIV persists in long-lived CD4⁺ T cell reservoirs despite suppressive ART, as integrated proviruses remain poised for reactivation. Chromatin remodeling is a central barrier to durable silencing, yet most studies have focused on SWI/SNF family members. The roles of non- SWI/SNF remodelers remain poorly defined, limiting our ability to rationally design host-directed “block-and-lock” cure strategies. Our unbiased shRNA screen of all 16 human remodeler ATPases identified EP400, CHD1, and CHD9 as repressors and INO80A, SMARCA5, and CHD2 as activators, establishing chromatin remodeling as a key determinant of HIV latency. Innovation: Our prior studies revealed that the p400 complex regulates HIV transcription through dual mechanisms: directly, by engaging Tat via the DMAP1 subunit to block Tat-TAR RNA interactions and restrict p-TEFb recruitment; and indirectly, by altering host transcriptional programs that control T cell activation states. Building on this mechanistic precedent and methodological platform, we now focus on INO80A, SMARCA5, CHD1, and CHD2, remodelers from distinct ATPase families that govern Tat-independent checkpoints at initiation, pause release, and elongation. Methodologically, we will apply TurboID-ChAP-MS (locus-specific proteomics), BEM-seq (single-nucleosome mapping), and degron-mediated acute depletion with ATPase-dead rescue to interrogate remodeler function with unprecedented resolution. Approach: Aim 1 will define the ATPase requirement and transcriptional checkpoints regulated by INO80A, SMARCA5, CHD1, and CHD2 using degron/CRISPR perturbations, ChIP-seq, nascent RNA profiling, and nucleosome mapping. Aim 2 will characterize remodeler-specific complexes and Tat dependence at the HIV promoter via TurboID proximity labeling integrated with chromatin affinity purification-mass spectrometry. Aim 3 will test combinatorial perturbations in Jurkat and primary CD4⁺ T cell latency models, including ART-suppressed donor cells, to identify synergistic “block-and-lock” strategies that enforce durable proviral silencing. Impact: By defining remodeler-specific mechanisms at discrete transcriptional checkpoints and leveraging their enzymatic, druggable activities, this work will establish chromatin remodeling as a therapeutic axis for durable HIV suppression and functional cure.

Molecular strategies for resolving differential regulation of dopamine subpopulations

Project Summary/Abstract Dopamine neurons in the ventral tegmental area (VTA) fire action potentials in complex patterns of tonic and phasic activity in response to environmental stimuli and during behavioral tasks. Transcriptomic, anatomical, and functional studies have established that VTA dopamine neurons can be divided into multiple subpopulations with variable gene expression, projection patterns, and response profiles. We recently completed a transcriptomic study that identified genetic markers for three distinct subpopulations of VTA dopamine neurons, and also found evidence for variability in ion channel gene expression between populations that correlated with differences in activity-dependent gene expression. However, much remains unknown regarding how specific genes encoding ion channels, receptors, transcription factors, or other signaling components contribute to the variability in baseline physiological properties observed across the VTA. Here we propose to combine slice electrophysiology recordings of VTA dopamine neurons with post-hoc single-cell sequencing analysis (i.e. patch-seq), which will allow us to directly correlate gene expression and physiological properties in order to identify candidate genes that may be key drivers of the variability between subpopulations. We also propose to validate and utilize a novel dual-recombinase CRISPR/Cas9 system for targeted gene mutagenesis in intersectional neuronal populations, which will provide a mechanism for testing gene function with unprecedented precision. We will use this approach to test the function of two candidate ion channel genes, the potassium channels Kcnh5 and Kcnh7, previously identified in our transcriptomic study as potential contributors to dopamine neuron action potential firing properties. We hypothesize that these genes are important for enabling rapid action potential firing in highly excitable dopamine neurons found in specific subpopulations. As a whole, with this proposal we aim to generate a valuable dataset linking gene expression in VTA dopamine neurons with physiology and subpopulation identification, as well as develop an intersectional gene mutagenesis strategy that can be used throughout the brain to precisely target neuronal subpopulations to test gene function. With this approach, we hope to facilitate future precision targeting of the dopamine system and dopamine-dependent behaviors.

Developing a novel technology for studying T cell differentiation in vivo

Summary CRISPR-based genetic screens have revolutionized our understanding of gene functions and molecular mechanisms across various biological processes. In the field of T cell biology, CRISPR screens have played a pivotal role in identifying genes that impact critical aspects, such as T cell development, differentiation, and function. However, traditional screens have struggled to distinguish genes with diverse mechanisms of action, necessitating further investigations. To address this challenge, researchers have harnessed the power of CRISPR screens combined with single-cell sequencing (scCRISPR-seq), enabling the simultaneous assessment of genetic perturbations and high-dimensional phenotypes at the single-cell level. While scCRISPR- seq has predominantly been performed in vitro using immortalized cell lines, its physiological relevance is limited due to oversimplified biological context and disparities compared to primary cells. This limitation highlights the urgent need for large-scale in vivo scCRISPR-seq with primary T cells. However, various challenges have discouraged its widespread adoption. The use of viral vectors for sgRNA delivery compromises physiological relevance, as the in vitro activation conditions fail to faithfully represent the intricate T cell priming process in vivo. Moreover, viral vector components and continuous Cas9 expression can trigger immunogenicity and cytotoxicity, leading to cell depletion and hindering long-term studies. Additionally, current scCRISPR-seq methods face technical limitations, including low editing efficiency and inadequate perturbation identity recovery rates, which impede efficient large-scale in vivo applications. Fortunately, recent advances in ribonucleoprotein complex (RNP) transfection have addressed many of these challenges. This cutting-edge technology enables efficient gene editing in primary T cells without the need for in vitro activation or permanent Cas9 expression. Leveraging the high editing efficiency of RNP transfection, the investigator’s team aims to develop a novel strategy for in vivo T cell CRISPR screens. This innovative approach involves arrayed RNP transfection and co- transfer of T cells that recognize the relevant antigens. Instead of traditional genetic barcodes, the strategy utilizes congenic markers (CD45.1/45.2 and CD90.1/CD90.2) from donor TCR transgenic T cells as "external barcodes." These markers facilitate the recovery of gene perturbation identity at the single-cell level through the application of CITE-seq. Importantly, this RNP-based strategy seamlessly integrates with existing single-cell sequencing protocols, enabling the comprehensive assessment of transcripts, epitopes, and chromatin accessibility simultaneously. To demonstrate the efficacy of this strategy, the team plans to develop two benchmarking approaches: RNP-CET-seq to investigate the role of TCR regulators in T cell exhaustion and RNP-CATE-seq to map the gene regulatory atlas of exhausted CD8 T cells. In summary, the proposed RNP- based scCRISPR-seq strategy overcomes the limitations of current approaches, enabling large-scale, multi- module in vivo genetic screens within a physiologically relevant context across various disease models.

Investigating the role of noncoding RNAs in malaria parasites through targeted Cas13-mediated degradation

Project Summary/Abstract One of the most significant sources of morbidity and mortality throughout large regions of the developing world continues to be malaria caused by infection with mosquito-borne parasites of the genus Plasmodium. The parasite species responsible for the most severe form of the disease is P. falciparum. To avoid antibodies produced by their host and thereby maintain lengthy infections, these parasites undergo a process called antigenic variation by which they can extend an infection for over a year. This results from changes in expression of a protein called PfEMP1, the primary antigenic and virulence determinant expressed on the surface of infected red blood cells. A large, multicopy gene family called var encodes different forms of PfEMP1, and switching expression between var genes enables parasites to evade antibody recognition and destruction by the immune system. The process requires precise and coordinated regulation of transcription of each var gene, however how this is accomplished is unknown. It was recently hypothesized that a family of noncoding RNAs (ncRNAs) plays a key role in controlling the expression of each var gene and in determining the likelihood of activation of any given gene. If correct, this would represent a significant advance in our understanding of how P. falciparum controls antigenic variation and avoids immune clearance. To test this hypothesis, we propose to adapt the CRISPR/Cas13 system of targeted RNA degradation for use in P. falciparum. Similar to the extensively used CRISPR/Cas9 system, CRISPR/Cas13 employes guide RNAs to target a nuclease to a sequence-specific target, however Cas13 targets single stranded RNA rather than DNA. By applying this system to the study of var-related ncRNAs, we will degrade specific ncRNAs and determine the effect on var gene expression. Two classes of ncRNAs previously proposed to regulate var gene expression will be targeted, one called ruf6 and a second encoded by the second exon of all var genes. This will enable us to alter ncRNA expression while leaving the underlying genomic DNA untouched, thereby allowing the unambiguous attribution of any resulting phenotypes to the ncRNAs. Aim 1 will optimize the Cas13 system for P. falciparum by testing different variants of the Cas13 endonuclease for their ability to degrade mRNAs encoding fluorescent reporter proteins. We will determine both the efficiency and sequence specificity of the system. Aim 2 will apply the system to var-associated ncRNAs and quantitatively measure changes in var gene expression and transcriptional switching. If successful, the adaptation of the Cas13 system to P. falciparum will provide the malaria research community with a powerful new tool for manipulating gene expression. In addition, we will gain valuable new insights into how malaria parasites regulate var gene expression, antigenic variation and immune evasion.

Engineering inducible morphotype switching control in Mycobacterium abscessus for investigating infection outcomes and discovering pathophysiological-targeted treatments

PROJECT SUMMARY Antibiotic-resistant nontuberculous mycobacteria (NTM) infections are rising at a rate of 8% each year and account for ~$1.7 billion in annual U.S. healthcare costs. Mycobacterium abscessus (Mabs), the most common rapidly growing NTM infection, is notoriously nicknamed the “antibiotic nightmare” for its extensive intrinsic and inducible broad-range multidrug resistance to antibiotic countermeasures. As part of its natural infection cycle, Mabs undergoes a morphotypical conversion from smooth to rough, characterized by irreversible genetic changes resulting in the loss of cell envelope glycopeptidolipids (GPLs). This morphotypic conversion is intimately associated with disease progression, ultimately leading to debilitating, refractory Mabs pulmonary disease. Specific stimuli triggering Mabs morphotypical conversion are unknown, thus preventing directed investigations into morphotype-specific immunological responses and the discovery of morphotype-specific therapeutic targets. This project leverages cutting-edge molecular genetic tools, including CRISPR (clustered regularly interspersed short palindromic repeats) interference (CRISPRi) and inducible knockdown control of CRISPRi via the anhydrotetracycline-inducible TetR-regulated promoter-operator system, to create six unique, reversible Mabs smooth to conditional rough morphotype strains. These molecular morphoswitchable strains allow precise investigator-mediated on-off control of Mabs surface GPLs, enabling investigations into Mabs morphological plasticity, unique pathophysiology traits associated with each morphotype, and the complex interplay between Mabs and morphotype-specific immunological responses. In Aim 1, we implement CRISPRi inducible knockdown tunable control of Mabs morphotype switching by targeting six, independent genetic targets directly involved in GPL biosynthesis (mps1, mps2) or transport (mmpS4, mmpL4a, mmpL4b, gap) and validate in vitro morphoswitching. In Aim 2, we establish and confirm Mabs morphoswitching and intracellular growth in infected THP-1 macrophages. Subsequently, we evaluate differential and distinct innate cellular immune responses elicited by Mabs smooth and Mabs conditional rough morphotypes during intracellular infection in human primary monocyte-derived macrophages. Collectively, these studies create a suite of characterized and reversible Mabs smooth and conditional rough morphoswitchable strains with controlled, regulated, and on- demand expression of Mabs surface GPLs. By enabling precisely timed and controlled induction of the Mabs conditional rough morphotype during intracellular growth, we can molecularly dissect and investigate fundamental Mabs host-pathogen interactions and immunological responses that so substantially influence negative clinical outcomes.

Towards Human Systems Biology of Sleep/Wake Cycles: Phosphorylation Hypothesis of Sleep

The field of human biology faces three major technological challenges. Firstly, the causation problem is difficult to address in humans compared to model animals. Secondly, the complexity problem arises due to the lack of a comprehensive cell atlas for the human body, despite its cellular composition. Lastly, the heterogeneity problem arises from significant variations in both genetic and environmental factors among individuals. To tackle these challenges, we have developed innovative approaches. These include 1) mammalian next-generation genetics, such as Triple CRISPR for knockout (KO) mice and ES mice for knock-in (KI) mice, which enables causation studies without traditional breeding methods; 2) whole-body/brain cell profiling techniques, such as CUBIC, to unravel the complexity of cellular composition; and 3) accurate and user-friendly technologies for measuring sleep and awake states, exemplified by ACCEL, to facilitate the monitoring of fundamental brain states in real-world settings and thus address heterogeneity in human.

Molecular recording using precision genome editing

SCN8A (Nav1.6) and DEE:  mouse models and pre-clinical therapies

SCN8A encodes a major voltage-gated sodium channel expressed in CNS and PNS neurons.  Gain-of-function and loss-of-function mutations contribute to  human disorders, most notably Developmental and Epileptic Encephalophy (DEE). More than 600 affected individuals have been reported, with the most common  mechanism of de novo, gain-of-function mutations.  We have developed constitutive  and conditional models of gain- and loss- of function mutations in the mouse and  characterized the effects of on neuronal firing and neurological phenotypes.  Using CRE lines with cellular and developmental specificity, we have probed the effects of activating  mutant alleles in various classes of neurons in the developing and adult mouse.   Most recently, we are testing genetic therapies that reduce the expression  of gain-of-function mutant alleles.  We are comparing the effectiveness of allele specific  oligos (ASOs), viral delivery of shRNAs, and allele-specific targeting of mutant alleles  using Crispr/Cas9 in mouse models of DEE.

Investigating activity-dependent processes in cerebral cortex development and disease

The cerebral cortex contains an extraordinary diversity of excitatory projection neuron (PN) and inhibitory interneurons (IN), wired together to form complex circuits. Spatiotemporally coordinated execution of intrinsic molecular programs by PNs and INs and activity-dependent processes, contribute to cortical development and cortical microcircuits formation. Alterations of these delicate processes have often been associated to neurological/neurodevelopmental disorders. However, despite the groundbreaking discovery that spontaneous activity in the embryonic brain can shape regional identities of distinct cortical territories, it is still unclear whether this early activity contributes to define subtype-specific neuronal fate as well as circuit assembly. In this study, we combined in utero genetic perturbations via CRISPR/Cas9 system and pharmacological inhibition of selected ion channels with RNA-sequencing and live imaging technologies to identify the activity-regulated processes controlling the development of different cortical PN classes, their wiring and the acquisition of subtype specific features. Moreover, we generated human induced pluripotent stem cells (iPSCs) form patients affected by a severe, rare and untreatable form of developmental epileptic encephalopathy. By differentiating cortical organoids form patient-derived iPSCs we create human models of early electrical alterations for studying molecular, structural and functional consequences of the genetic mutations during cortical development. Our ultimate goal is to define the activity-conditioned processes that physiologically occur during the development of cortical circuits, to identify novel therapeutical paths to address the pathological consequences of neonatal epilepsies.

The history, future and ethics of self-experimentation

Modern day “neurohackers” are radically self-experimenting, attempting genomic modification with CRISPR-Cas9 constructs and electrode insertion into their cortex amongst a host of other things. Institutions wanting to avoid the risks bought on by these procedures, generally avoid involvement with self-experimenting research. Modern day “neurohackers” are radically self-experimenting, attempting genomic modification with CRISPR-Cas9 constructs and electrode insertion into their cortex amongst a host of other things. Institutions wanting to avoid the risks bought on by these procedures, generally avoid involvement with self-experimenting research. But what is the ethical thing to do? Should researchers be allowed or encouraged to self-experiment? Should institutions support or hinder them? Where do you think that this process of self-experimentation could take us? This presentation by Dr Matt Lennon and Professor Zoltan Molnar of the University of Oxford, will explore the history, future and ethics of self-experimentation. It will explore notable examples of self-experimenters including Isaac Newton, Angelo Ruffini and Oliver Sacks and how a number of these pivotal experiments created paradigm shifts in neuroscience. The presentation will open up a forum for all participants to be involved asking key ethical questions around what should and should not be allowed in self-experimentation research.

Towards targeted therapies for the treatment of Dravet Syndrome

Dravet syndrome is a severe epileptic encephalopathy that begins during the first year of life and leads to severe cognitive and social interaction deficits. It is mostly caused by heterozygous loss-of-function mutations in the SCN1A gene, which encodes for the alpha-subunit of the voltage-gated sodium channel (Nav1.1) and is responsible mainly of GABAergic interneuron excitability. While different therapies based on the upregulation of the healthy allele of the gene are being developed, the dynamics of reversibility of the pathology are still unclear. In fact, whether and to which extent the pathology is reversible after symptom onset and if it is sufficient to ensure physiological levels of Scn1a during a specific critical period of time are open questions in the field and their answers are required for proper development of effective therapies. We generated a novel Scn1a conditional knock-in mouse model (Scn1aSTOP) in which the endogenous Scn1a gene is silenced by the insertion of a floxed STOP cassette in an intron of Scn1a gene; upon Cre recombinase expression, the STOP cassette is removed, and the mutant allele can be reconstituted as a functional Scn1a allele. In this model we can reactivate the expression of Scn1a exactly in the neuronal subtypes in which it is expressed and at its physiological level. Those aspects are crucial to obtain a final answer on the reversibility of DS after symptom onset. We exploited this model to demonstrate that global brain re-expression of the Scn1a gene when symptoms are already developed (P30) led to a complete rescue of both spontaneous and thermic inducible seizures and amelioration of behavioral abnormalities characteristic of this model. We also highlighted dramatic gene expression alterations associated with astrogliosis and inflammation that, accordingly, were rescued by Scn1a gene expression normalization at P30. Moreover, employing a conditional knock-out mouse model of DS we reported that ensuring physiological levels of Scn1a during the critical period of symptom appearance (until P30) is not sufficient to prevent the DS, conversely, mice start to die of SUDEP and develop spontaneous seizures. These results offer promising insights in the reversibility of DS and can help to accelerate therapeutic translation, providing important information on the timing for gene therapy delivery to Dravet patients.

Harnessing the CRISPR toolbox to engineer biology

Sparks, flames, and inferno: epileptogenesis in the glioblastoma microenvironment

Glioblastoma cells trigger pharmacoresistant seizures that may promote tumor growth and diminish the quality of remaining life. To define the relationship between growth of glial tumors and their neuronal microenvironment, and to identify genomic biomarkers and mechanisms that may point to better prognosis and treatment of drug resistant epilepsy in brain cancer, we are analyzing a new generation of genetically defined CRISPR/in utero electroporation inborn glioblastoma (GBM) tumor models engineered in mice. The molecular pathophysiology of glioblastoma cells and surrounding neurons and untransformed astrocytes are compared at serial stages of tumor development. Initial studies reveal that epileptiform EEG spiking is a very early and reliable preclinical signature of GBM expansion in these mice, followed by rapidly progressive seizures and death within weeks. FACS-sorted transcriptomic analysis of cortical astrocytes reveals the expansion of a subgroup enriched in pro-synaptogenic genes that may drive hyperexcitability, a novel mechanism of epileptogenesis. Using a prototypical GBM IUE model, we systematically define and correlate the earliest appearance of cortical hyperexcitability with progressive cortical tumor cell invasion, including spontaneous episodes of spreading cortical depolarization, innate inflammation, and xCT upregulation in the peritumoral microenvironment. Blocking this glutamate exporter reduces seizure load. We show that the host genome contributes to seizure risk by generating tumors in a monogenic deletion strain (MapT/tau -/-) that raises cortical seizure threshold. We also show that the tumor variant profile determines epilepsy risk. Our genetic dissection approach sets the stage to broadly explore the developmental biology of personalized tumor/host interactions in mice engineered with novel human tumor mutations in specified glial cell lineages.

CRISPR-based functional genomics in iPSC-based models of brain disease

Human genes associated with brain-related diseases are being discovered at an accelerating pace. A major challenge is an identification of the mechanisms through which these genes act, and of potential therapeutic strategies. To elucidate such mechanisms in human cells, we established a CRISPR-based platform for genetic screening in human iPSC-derived neurons, astrocytes and microglia. Our approach relies on CRISPR interference (CRISPRi) and CRISPR activation (CRISPRa), in which a catalytically dead version of the bacterial Cas9 protein recruits transcriptional repressors or activators, respectively, to endogenous genes to control their expression, as directed by a small guide RNA (sgRNA). Complex libraries of sgRNAs enable us to conduct genome-wide or focused loss-of-function and gain-of-function screens. Such screens uncover molecular players for phenotypes based on survival, stress resistance, fluorescent phenotypes, high-content imaging and single-cell RNA-Seq. To uncover disease mechanisms and therapeutic targets, we are conducting genetic modifier screens for disease-relevant cellular phenotypes in patient-derived neurons and glia with familial mutations and isogenic controls. In a genome-wide screen, we have uncovered genes that modulate the formation of disease-associated aggregates of tau in neurons with a tauopathy-linked mutation (MAPT V337M). CRISPRi/a can also be used to model and functionally evaluate disease-associated changes in gene expression, such as those caused by eQTLs, haploinsufficiency, or disease states of brain cells. We will discuss an application to Alzheimer’s Disease-associated genes in microglia.

Creating cholinergic neuron specific knock-out mice by combining three (CRISPR-Cas9, Cre/loxP and AAV) genome editing technologies

CRISPR-based epigenetic editing of engram cells in fear memories

CRISPR/Cas9 gene-edited fluorescent organoids as a novel model to study Alzheimer’s disease pathology

CRISPR-Mediated Activation of Autism Gene Itgb3 Restores Cortical Network Excitability via mGluR5 Signaling

Developing zebrafish CRISPR/Cas9 knockout models of Parkinson’s Disease to identify novel therapeutic targets

The effect of a GRIN2D variant in Developmental and Epileptic Encephalopathy in a CRISPR/Cas9 mouse model and iPS cells

Whole-genome CRISPR interference screen identifies ARID1A-dependent growth regulators in human induced pluripotent stem cells

Application of single-cell CRISPRi/a screen to characterize multiple sclerosis-associated single nucleotide polymorphisms in oligodendrocytes

FENS Forum 2024

CRISPR-based epigenetic editing of engram cells in fear memories

FENS Forum 2024

CRISPR-mediated approaches for ATXN2 knockdown in SCA2 models of disease

FENS Forum 2024

Establishing an experimental sgRNA expression screening assay for CRISPR activation in vitro

FENS Forum 2024

Establishment of animal cell lines to detect circadian rhythm gene expression via CRISPR-Cas9

FENS Forum 2024

Single-cell CRISPR screening in cerebral organoids identifies developmental and cell type-specific defects of autism

FENS Forum 2024

Understanding CaV2.1 dysfunction in neurological disorders: Insights from novel CRISPR/Cas9 mouse model and iPSC-derived neurons

FENS Forum 2024