Multinucleated giant cells (MGCs) are a hallmark pathological feature of a wide range of diseases, yet the mechanisms underlying their formation and function remain poorly understood. Although it is recognized that MGCs arise from a heterogeneous pool of myeloid precursors, how a committed MGC fate develops therein remains unknown. Here, combining temporal in vitro and in vivo differentiation of bone marrow-derived myeloid precursors with single cell and bulk RNA sequencing as well as CRISPR/Cas9-mediated gene editing, we shed insight into how MGCs emerge. Our findings reveal that coordinated upregulation of cell fusion genes and cellular metabolism, particularly oxidative phosphorylation, regulate the transition from macrophage progenitors to fusion competent cells. In vivo fate-mapping unveils Ms4a3 -, and Cd11c -but not Cx3cr1 -traced cells as the predominant precursor populations for MGCs in a lung granuloma formation model. Notably, transcription factor profiling in the progression from early myeloid precursors to pre-MGCs identifies IRF4 as a key molecular switch driving MGC generation. IRF4 + MGCs are present in different pathologies, including Schistosoma mansoni egg induced granulomas, Aspergillus fumigatus conidia mediated allergic airway inflammation and human head and neck squamous cell carcinomas. Mechanistically, IRF4 controls critical fusion related genes such as Dcstamp and Ocstamp . Consequently, Irf4 deficient cells are unable to develop into MGCs. Collectively, our work delineates the trajectory of MGC differentiation, establishing IRF4 as a defining transcription factor required for the generation of fusion-competent progenitors that ultimately give rise to MGCs.

CRISPR gene editing has revolutionized our ability to study and manipulate specific genes, enabling novel insights into gene function and potential therapies for brain disorders. Recent advances in cell-type-specific regulatory elements and viral delivery systems have made precise in vivo gene editing possible. However, neurons with similar molecular profiles can belong to different circuits, complicating efforts to manipulate circuit function and behavior. To address this, we developed CRISPR-rabies virus (CRV), which leverages the trans-synaptic spread of rabies virus to enable gene editing within anatomically defined neural circuits. By pairing CRV with cell type-specific Cas9 expression, we achieved targeted gene modifications in specific circuits. We demonstrate that CRV can modulate sodium and potassium channel expression in parvalbumin interneurons, thereby effectively regulating synaptic transmission of pyramidal neurons in the CA3 region of the hippocampus. Its compatibility with 3′-capture single-cell RNA-seq allows simultaneous circuit perturbation and molecular profiling. In summary, CRV allows precise circuit-level gene modulation, providing a platform for studying gene function in neural circuits and developing novel gene therapies for brain disorders.

The type I-E and I-F CRISPR-Cas systems were identified in 237 E. coli strains isolated from patients with urinary tract infections (UTIs) between 2004 and 2019. The strains were classified into nine distinct groups (I-IX) based on the presence or absence of cas genes and repeat regions (RRs). Within the type I-E systems, two sequence variants were identified, distinguished by polymorphisms in the casB, cas3, cas7, cas5, and cas6 genes. The direct repeats (DRs) also differed, with I-E-associated RRs ranging from 26–32 bp and I-F-associated RRs being a consistent 28 bp. We identified 762 unique spacers (29–35 bp in length) across the strain collection. The number of spacers per strain varied from 1 to 47, and potential DNA targets were determined for 83 spacers, targeting 56 bacteriophage genomes, 19 plasmids, and 8 cas genes of the I-F type. Multilocus sequence typing (MLST) revealed 68 sequence types and 24 clonal complexes (CCs), with ST131, CC10, CC69, CC405, CC14, CC38, CC73, and CC648 being the most prevalent. Significant correlations were observed between specific phylogroups/CCs, the type of CRISPR-Cas system present, and distinct profiles of virulence and antibiotic resistance genes.

The selective loss of dopaminergic neurons in the substantia nigra is a hallmark of Parkinson’s disease (PD), yet the contribution of glial cells to this vulnerability is not fully understood. Studies in rodent models suggest that astrocytes can take up and metabolize dopamine (DA), potentially protecting neurons by detoxifying reactive DA metabolites via glutathione S-transferase mu 2 (GSTM2) release. However, these mechanisms remain underexplored in human systems, particularly in the context of PD. Here, we used CRISPR-engineered iPSC-derived human astrocytes with a PD-linked DJ-1 mutation and isogenic controls to investigate astrocytic DA metabolism. Upon DA exposure, control astrocytes upregulated quinone-reducing enzymes NAD(P)H quinone dehydrogenase 1 (NQO1) and GSTM2, whereas DJ-1 mutant astrocytes failed to adaptively respond. In addition, only control astrocytes presented with increased DA quinone products upon DA exposure, not DJ-1 mutants. These results demonstrate astrocytic DA handling being disrupted in DJ-1-linked PD, implicating astroglial dysfunction as an important contributor to PD pathogenesis and potential target for therapeutic intervention.

Summary ER-phagy receptors have elusive physiological functions beyond ER remodelling. Here, we use proximity biotinylation to identify their cytoplasmic interactomes. Secondary CRISPR/Cas9 screening reveals regulators of the prototypical FAM134B/C receptors, which include PRKAR1A, canonically known as a subunit of PKA. PRKAR1A directly binds an amphipathic helix in the otherwise disordered cytoplasmic domain of FAM134B. Super-resolution, FRAP and CLEM imaging reveal novel interorganellar contacts between liquid-like condensates of cAMP-bound PRKAR1A and the ER. Condensates promote clustering of ER-embedded FAM134B/C with LC3B, independently of regulation of PKA by PRKAR1A. Proteomics reveal that cytoplasmic RhoA interacts with FAM134B/C clusters and that sequestration of both these molecules occurs within lysosomes embedded within the proximal condensates. This results in reduced actomyosin contractility and ER-condensate interactions thusly determine cell morphology and cancer cell invasion modality. In summary, ER-condensate contacts mediated by FAM134B/C are novel cellular degradation hubs that coordinate ER and cellular remodelling.

From mammals to bacteria, the direct recognition and cleavage of viral nucleic acids is a potent defense strategy against viral infection, but it requires mechanisms for distinguishing self from non-self 1,2 . In bacteria, CRISPR-Cas and restriction modification systems achieve this discrimination by recognizing specific DNA sequences or DNA modifications. Alternative mechanisms likely remain to be discovered. Here, we characterize SNIPE, a novel anti-phage defense system that constitutively localizes to the bacterial cell membrane in E. coli to block phage λ infection. Using radiolabeled phage DNA and time-lapse microscopy to track phage genomes we demonstrate that SNIPE directly cleaves phage DNA during genome injection. Based on proximity labeling, we find that SNIPE associates with host proteins essential for λ genome entry and with the λ tape measure protein, which facilitates λ genome injection across the inner membrane. SNIPE also defends against diverse siphoviruses, likely through direct interactions with their tape measure proteins. Our findings establish SNIPE as a widespread bacterial defense system that exploits the spatial organization of phage genome injection to specifically target viral DNA, representing a novel strategy for distinguishing self from non-self in prokaryotic immune systems.

ABSTRACT Type I IFN, including IFN-α, induces the expression of antiviral restriction factors that can interfere with multiple steps of the HIV-1 replication cycle. Humans have 13 IFN-α genes which encode 12 different IFN-α subtypes. Our previous work in HIV-1 infected humanized mice showed that IFN-α14 treatment more potently controlled HIV-1 than treatment with the clinically approved IFN-α2 subtype. However, the mechanisms behind the more potent control of HIV-1 by IFN-α14 are unknown. The IFN-α14 subtype is known to more potently induce the expression of the restriction factors MX2 and ISG15 and increased APOBEC3G signature mutations in vivo compared to IFN-α2. To study the importance of each of these restriction factors in mediating the potent control of HIV-1, we used a CRISPR-Cas9 lentivirus system to create stable knockouts in the MT4C5 cell line that is susceptible to HIV-1 but does not produce measurable amounts of endogenous IFN-α. Knock out of ISG15, but not MX2, eliminated differences in viral suppression after IFN-α14 and IFN-α2 treatment. Similarly, APOBEC3G deletion eliminated differences in viral suppression and the number of infectious particles produced after IFN-α14 and IFN-α2 treatment. Furthermore, APOBEC3G deletion resulted in significantly fewer GG→AG mutations in viral DNA isolated from target cells incubated with supernatant from IFN-α14 treated groups. However, APOBEC3G knock out did not result in significant increases in vDNA compared to the wild type in any experimental group. Overall, elimination of APOBEC3G and ISG15 impaired IFN-α14–mediated suppression of HIV-1, highlighting them as downstream effectors of IFN-α14’s more potent anti-HIV-1 activity. IMPORTANCE This study uncovers the molecular basis for the more potent antiviral activity of IFN-α14 compared to the clinically used IFN-α2 subtype against HIV-1. Although interferons are known to induce numerous restriction factors, the mechanisms underlying subtype-specific antiviral potency remained unclear. By using CRISPR-Cas9 knockout MT4C5 cell lines, the study identifies ISG15 and APOBEC3G as key effectors mediating IFN-α14’s enhanced suppression of HIV-1 replication. Loss of either ISG15 or APOBEC3G abolished the differential antiviral effect between IFN-α14 and IFN-α2, demonstrating their essential roles in IFN-α14 driven viral restriction. These findings highlight that individual IFN-α subtypes engage distinct downstream pathways and that subtype diversity encodes functional specialization rather than redundancy. Overall, this work advances our understanding of innate immune control of HIV-1 and provides a foundation for developing targeted interferon-based therapies that exploit the unique mechanisms of potent subtypes like IFN- α14.

The engineering of autologous T cells for the expression of chimeric antigen receptors (CARs) can induce profound clinical responses in haematological malignancies, while T cell receptor-engineered T (TCR T) cells have led to durable responses to solid tumours in clinical trials. However, the clinical production of engineered T cells is exhaustive and often leads to highly differentiated and exhausted effector T cells. To circumvent this, we have developed an antigen-scaffold (Ag-scaffold) technology to preferentially expand genetically engineered T cells. Such Ag-scaffolds present cognate antigen together with stimulatory factors such as cytokines. By providing a specific and receptor-engaging stimulation to CAR/TCR-engineered T cells, the expanded product is highly enriched for engineered T cells with a favourable proliferative and efficacious phenotype. Here, we expand CRISPR/Cas9- and lentiviral-engineered TCR T and CAR T cells. We expanded TCR T-cells with Ag-scaffolds presenting peptide MHC (pMHC), and anti-CD19 CAR T cells with Ag-scaffolds presenting CD19 antigen. By applying cognate pMHC Ag-scaffolds, we achieved more than 90% antigen-specific T cells after 14 days of culture with a distinct cytotoxic, proliferative phenotypical profile. Ag-scaffold expansion enhanced initial TCR and CAR cytotoxicity; sustained control was observed after repeated rechallenges of CAR T cells. In vivo , Ag-scaffold-expanded CRISPR/Cas9-engineered anti-CD19 CAR T also showed complete tumour eradication in a B-cell lymphoma xenograft model with a low dose of CAR T cells, which was not achieved using IL2/7/15 expansion.

ABSTRACT Several studies examined host and pathogen genetic influences on tuberculosis (TB) susceptibility separately, but relatively few studied their combined effects. However, host-pathogen interactions or co-evolution may explain the inability to replicate many reported human genetic effects across global populations and provide additional insight into TB risk. In this study, we address such possible interactions by focusing on the outcome of infection with L4-Uganda M. tuberculosis sub-lineage and human genetic variants as independent variables. This is possible because the L4-Uganda sub-lineage is both restricted to Uganda and nearby locations and is recent there, compared to other more ancestral L4 lineages. Our study consisted of 276 culture-confirmed adult TB cases from a long-standing household contact study. Multiple loci with results suggestive of association (p<10 -5 ) also demonstrated convergent relevant evidence for strain specific infection via: evidence of gene expression in relevant cells and lung tissue, signatures of natural selection, eQTL expression, and CRISPR screens for immunity-related genes. We also replicated previously published host-pathogen interaction effects, demonstrating that effects seen for other sub-lineages were also present for L4-Uganda. These results provide evidence for host-pathogen co-evolution in TB and indicate these interactions involve genes highly relevant to the host immune response to Mycobacterium infection.

Summary Mutations in the mitochondrial DNA (mtDNA) are associated with severe human diseases, lacking efficient therapies. Direct correction of mtDNA mutations may offer a cure for such diseases. We propose a novel strategy based on double-stranded DNA (dsDNA) oligonucleotide delivery into mitochondria and intrinsic microhomology-mediated end joining (MMEJ) for mtDNA editing. This strategy enables introduction of multiple predefined nucleotide changes in mtDNA. For this, the presence of MMEJ activity in the human mitochondrial lysates was confirmed. 49 bp DNA oligonucleotide duplexes, fused to an RNA hairpin previously identified as a mitochondrial import signal, were delivered into the mitochondria of cultured human cells. Delivery of these donor dsDNA molecules, homological to an ND4 site of mtDNA and bearing designed nucleotide changes, led to a low but statistically significant introduction of the designed nucleotide changes into mtDNA. Donor dsDNA delivery combined with the CRISPR/mito-AsCas12a system also resulted in a statistically significant number of an expected concomitant change of five nucleotides distributed across a 16-nucleotide ND4 site of the mitochondrial genome. The proposed strategy may become an efficient mtDNA editing tool suitable for the correction of near-homoplasmic mutations such as Leber’s Hereditary Optic Neuropathy (LHON)-associated mutations in the ND4 gene of mtDNA.

CRISPR-associated transposons (CAST) use guide RNAs to direct their transposition and are being harnessed as tools for programmable genome engineering across diverse bacterial species. However, CAST systems have not been adapted for high-throughput genetic screening. Here, we present MultiCAST, a streamlined platform for rapid and scalable guide RNA-directed transposon insertion in bacteria. MultiCAST generates targeted insertions in a single step through conjugative delivery of conditionally replicative plasmids encoding the CAST enzymatic machinery and a selectable mini-transposon expressing the guide RNA. By leveraging the inserted guide sequence as a molecular barcode, MultiCAST enables pooled, high-throughput genetic screens using only amplicon sequencing. We identified factors that influence transposition efficiency and the accuracy of insertion frequency measurements derived from guide sequencing. Adjusting the ratio of donor and recipient strain during conjugation mitigates guide-transposon “crosstalk”, in which a single recipient cell acquires multiple donor plasmids containing distinct guides. Furthermore, we developed a machine learning-based predictive model for selecting highly active guides based on target sequence features that strongly correlate with activity. The nucleoid-associated protein H-NS was also found to inhibit CAST activity, providing a mechanistic explanation for variable insertion frequencies among non-essential genes. To demonstrate the scalability of MultiCAST, we screened a pooled mutant population created from >5,200 guides targeting 88 genes in E. coli across twelve nutrient conditions, accurately identifying genes with condition-specific fitness effects. The simplicity, speed, and throughput of MultiCAST make genome-scale functional screens more accessible across a wide range of bacterial species. Significance Efficient gene disruption is essential for understanding bacterial gene function, but traditional genetic approaches are labor-intensive and generally not well-suited for high-throughput studies. We developed MultiCAST, a simple and scalable method that harnesses guide RNA-directed CRISPR-associated transposons for targeted bacterial gene disruption. MultiCAST enables single and pooled transposon mutagenesis in a single step and eliminates the need for complex sequencing library preparation protocols by using the guide sequence as a quantifiable surrogate for mutant abundance. This approach allows thousands of mutants to be generated and screened simultaneously across multiple conditions using only amplicon sequencing. By dramatically reducing the time, cost, and complexity of reverse genetics, MultiCAST opens new possibilities for genome-scale functional studies, accelerating the discovery of bacterial gene functions.

Streptococcus pneumoniae is an important human pathogen that causes invasive diseases and remains the leading cause of bacterial meningitis. However, the bacterial determinants required for survival within the central nervous system (CNS) are poorly understood. Here, we performed a genome-wide CRISPR interference screen (CRISPRi-Seq) in an in vivo zebrafish model to systematically map pneumococcal fitness determinants during meningitis. Using this approach, we investigated essential and non-essential genes contributing to the pathogenesis of pneumococcal meningitis. The screen identified 244 loci whose repression significantly reduced bacterial fitness in vivo , representing pathways involved in virulence, metabolism, cell-envelope biogenesis, translation, and stress adaptation. Comparative analysis with in vitro datasets showed that metabolic genes such as purA, proABC, thrC, glyA, and manLMN, as well as those involved in peptidoglycan and capsule biosynthesis (pbp3, cps2 operon) and oxidative stress response ( nox, dpr ), become selectively more essential in vivo. Functional validation confirmed that these pathways are critical for virulence, nutrient-dependent growth, and oxidative stress response during meningitis. Untargeted metabolomics of infected zebrafish corroborate several observed bacterial genetic sensitivities in the CNS with altered levels of key nutrients such as glucose and 4-aminobenzoic acid. Integration of the CRISPRi-Seq dataset with antibiotic-target annotations revealed both established and previously unrecognised vulnerabilities, including aminoacyl-tRNA synthetases such as leucyl-tRNA synthetase (LeuRS), whose inhibition by the benzoxaborole epetraborole improved host survival, highlighting its potential as a new therapeutic strategy for multidrug-resistant (MDR) S. pneumoniae. Together, these findings demonstrate the power of in vivo CRISPRi-Seq to define the pneumococcal essentialome under physiological conditions and reveal that metabolic adaptation, cell-envelope maintenance, and the oxidative stress response are central to bacterial survival in the CNS. This fitness map advances our understanding of pneumococcal adaptation in the CNS and identifies promising targets for developing therapies against S. pneumoniae .

Toxoplasma gondii undergoes sexual development exclusively in the feline intestine, a process critical for genetic diversity and population expansion. Recent studies have identified genes critical in suppressing presexual development 1 and metabolic differences in felines that may promote sexual development 2 , but to date the gene regulatory networks driving development in the cat are unknown. To investigate this, we performed single-cell transcriptomics on parasites isolated from cat intestines, using fluorescent reporter strains and flow cytometry. From 15,068 cells across two experiments, we identified rare populations, including cells that bear all of the hallmarks of gametes. Candidate genes emerging from this study were tested via CRISPR-Cas9 Perturb-seq, identifying AP2X6 as a regulator of macrogametocyte development. Our single-cell data extends what is known about gene expression changes throughout sexual development and should be useful to those in the field working towards inducing gametogenesis, mating, and oocyst production in vitro .

Fluorescent in situ sequencing involves imaging-based sequencing by synthesis in intact cells or tissues to reveal target nucleotide sequences inside each cell. Often, the target sequences are barcodes that indicate a perturbation (e.g. CRISPR guide or genetic variant) delivered to the cell. However, processing in situ sequencing data presents a considerable challenge, requiring stitching and aligning tens of thousands of images with millions of cells, detecting small amplicon colonies across sequencing cycles, and calling reads. To address these challenges, we introduce STARCall: STitching, Alignment and Read Calling for in situ sequencing, a software package that analyzes raw in situ sequencing images to produce a genotype-to-phenotype mapping for each cell. STAR-Call improves upon previous solutions by combining stitching and alignment of images into a single step that minimizes both inter-cycle and intra-cycle alignment error. STARCall also improves detection and extraction of sequencing reads, incorporating filters and normalization to combat background fluorophore signal. We compare STARCall to other methods using a diverse set of images that include commonly encountered imaging problems such as variable intensity across channels and cycles and high levels of background. Specifically, this comprises ∼250,000 images from a pooled screen of ∼3,500 barcoded LMNA variants expressed in U2OS cells and ∼1,200 bar-coded PTEN variants in induced pluripotent stem cells (iPSC) and iPSC-derived neurons. Overall, STARCall aligned more than 50% of tiles with <1 pixel error on all nine image sets while alternative packages had higher error on four. STARCall also yielded an 8-40% increase in genotyped cells due to improved filtering and normalization methods that address background fluorescence. STARCall can call tools like CellPose to segment cells and CellProfiler to compute cell features from the phenotyping images. STARcall is open-source and freely available, providing a robust solution for the analysis of in situ sequencing data. Author summary Short regions of RNA or DNA can be sequenced inside intact cells or tissues (i.e. in situ ) by combining a microscope and sequencing by DNA synthesis. Multiple cycles of sequencing are performed, in which incorporation of a single fluorescently labeled nucleotide is imaged, and the corresponding base detected. Recently, in situ sequencing has proved useful in optical pooled screens, where a library of perturbations such as CRISPR-mediated gene knockouts or genetic variants is introduced into cells, and in situ sequencing is used to reveal the specific perturbation in each cell. However, processing in situ sequencing data presents a considerable challenge, requiring stitching and aligning tens of thousands of images, detecting small amplicon colonies across sequencing cycles, and calling reads. To address these challenges, we introduce STARCall: STitching, Alignment and Read Calling for in situ sequencing. STARCall uses a novel stitching algorithm that minimizes both inter-cycle and intra-cycle alignment error, and improved filters and normalization for base calling. When applied to a set of 9 in situ sequencing image sets, STARCall yielded an 8-40% increase in genotyped cells. STARCall is open-source and applicable to a variety of experiments, providing a robust pipeline for in situ sequencing data.

ABSTRACT Diatoms are globally significant microalgae that contribute ∼ 20% of oxygen production and exhibit remarkable metabolic diversity. The marine diatom Phaeodactylum tricornutum has emerged as a promising synthetic biology platform for bioproduction of recombinant proteins, supported by a human-like N -linked glycosylation pathway. However, its α (1,3)-linked core fucose is immunogenic in humans and thus limits biopharmaceutical applications. One hurdle to efficient genome engineering in P. tricornutum is the lack of a robust system for simultaneous CRISPR/Cas9 editing at multiple sites. To overcome this limitation, we develop PHYCUT ( Ph aeodactylum tricornutum Cs y 4- C as9 m u ltiplex t ool), a versatile plasmid-based CRISPR/Cas9 system that uses the Csy4 endoribonuclease to process multi-guide RNA arrays. To highlight PHYCUT applications, we demonstrate multiplex editing of all three FucT genes responsible for α (1,3) fucosylation in P. tricornutum , yielding strains with markedly reduced fucosylation of secreted proteins. PHYCUT enables facile, multiplexed genome engineering in diatoms and provides a foundation for humanizing the P. tricornutum glycosylation pathway to support next-generation algal biotechnology.

Bacteriophage anti-CRISPR (Acr) proteins have the potential to reduce off-target effects of genome editing by inactivating the CRISPR-Cas bacterial defense. The current challenge lays in their functional annotation, as Acr proteins have high structural diversity and low sequence similarity, thus rendering common homology-based methods unfit. Recent solutions use deep learning models such as graph convolutional networks that take protein networks as the data input. In an effort to understand whether these new solutions are fit for niche, sparsely annotated proteins, we focus on 3 Acr proteins (AcrIF1, AcrIIA1, and AcrVIA1) as a case study. For each, we create protein contact networks (PCNs) and residue interaction graphs (RIGs) based on existing network theory and methodology. We characterize and analyze these protein networks by comparing how each network architecture affects values of small-worldliness. We reexamine a previous method that focused on using node degree, closeness centralities, and residue solvent accessibility to predict functional residues within a protein via a Jackknife technique. We discuss the implications of the construction of these networks based on how the structure information is acquired. We demonstrate that functional residues within small proteins cannot be reliably predicted with the Jackknife technique, even when provided with a curated dataset containing representative standardized values for degree and closeness centrality. We show that functional residues within these small proteins have low degrees within both PCNs and RIGs, thus making them susceptible to the known degree bias towards high degree nodes present in using graph convolutional networks. We discuss how understanding the data can be used to further improve deep learning approaches for small proteins. Author summary A bacteria’s CRISPR-Cas defense system acts as security guard against viruses like bacteriophages. By storing pieces of viral DNA as records, it can recognize and defend the bacteria against threats. Scientists have adapted this effective record keeping process to perform targeted genome editing. Some bacteriophages have genes that encode for anti-CRISPR (Acr) proteins. The proteins act as a criminal accomplice to the viral DNA, sneaking them in past the bacteria’s security in a variety of ways. There has been increased interest in using these Acr proteins to limit unintended or off-target effects of targeted genome editing. However, Acr proteins are difficult to identify. We changed parts of a previous method that used graph representations of protein structure to determine important amino acids that help that protein perform its function. We applied these methods to three Acr proteins to determine whether we observed similar patterns in these graphs. We explain how features of these graph representations of protein structures can affect graph neural networks that use them as input to learn more about proteins.

Urinary tract infections (UTIs) are among the most common infectious diseases, causing over 400 million cases and 260,000 deaths annually. Women are disproportionately affected, with ∼50% experiencing at least one UTI during their lifetime and 20–30% suffering from recurrent infections. Uropathogenic Escherichia coli (UPEC), which accounts for ∼75% of cases, employs diverse virulence factors to persist and evade host immunity. Rising antibiotic resistance, driven by widespread antimicrobial misuse, is eroding treatment efficacy and highlights the urgent need for alternative therapeutic strategies. To uncover novel vulnerabilities under physiologically relevant conditions, we constructed a genome-wide CRISPR interference (CRISPRi) library in the UPEC reference strain E. coli CFT073 and systematically profiled gene fitness in rich media versus human urine. The screen revealed multiple pathways that are conditionally essential for UPEC growth in urine, including iron uptake, envelope maintenance, and the biosynthesis of arginine, methionine, and branched-chain amino acids. Notably, we identified acetolactate synthase (ALS) II as the sole active isoform supporting branched-chain amino acid synthesis in urine. Functional validation further demonstrated its druggability: introducing a re-sensitizing mutation overcame the protein’s intrinsic resistance to the ALS-targeting herbicide sulfometuron methyl, restoring sensitivity. These findings establish ALS II as a promising therapeutic target against UPEC.

The spring barley cultivar Golden Promise (GP) is the major reference genotype for transformation due to its high transformability and availability of a reference genome. However, GP is characterized by a long generation cycle and stress susceptibility under non-optimal growth conditions because it carries a mutation at the floral inducer Photoperiod-H1 ( Ppd-H1 ). Previously, we showed that a GP introgression line, Golden Promise-fast (GP-fast), generated by introducing the wild-type Ppd-H1 allele from the winter barley cultivar Igri, exhibits early flowering and improved stress resilience. In this study, we generated a fast-cycling genotype, Golden Promise-rapid (GP-rapid), isogenic to GP with high transformation efficiency. We conducted two backcrosses of GP-fast to reduce the residual Igri genome. The resulting genotype contains only a single introgression of approximately 0.6 Mbp at the Ppd-H1 locus on chromosome 2H. Under speed breeding conditions, its generation time was reduced to 63 days (25% shorter than GP’s 84 days). Parallel transformation of GP, GP-fast, and GP-rapid using CRISPR/Cas9-mediated genome editing of Ppd-H1 revealed high regeneration and transformation efficiencies of GP-rapid, comparable to GP. Overall, we report on the development of a fast-cycling GP isogenic line as a research tool for efficient generation of transgenic and gene-edited barley plants. Highlights A new fast-cycling barley genotype, GP-rapid, reduces generation time by 25% while retaining high transformation efficiency, advancing functional genomic studies in barley.

Abstract Obesity and neurodegeneration are clinically associated diseases with defective autophagy. However, the genetic, biological, and metabolic underpinnings connecting these diseases are not well-understood. Here we identified a Mitochondria obesity/neurodegeneration (M on ) gene-signature that is shared between obesity, and neurodegenerative diseases. We demonstrate that, CEBPB elevates M on -gene-signature, to form podosomal belts, and enhance ROS production. Inhibiting autophagy collapses podosomal-belts through macropinocytosis to accumulate vacuoles, lipid-droplets, nuclear Notch-1 (nNICD), DEPTOR, and HBV-polymerase mRNAs. Conversely, hemin counteracts these events and suppresses DEPTOR and HBV-polymerase mRNAs by A-to-I-RNA-editing and nonsense-mediated decay. Furthermore, we CRISPR-engineered the antiviral chromosome-19 miRNA cluster (C19MC) to demonstrate that C19MC-miRNAs augment CEBPB, M on -gene-signature, ROS, and recapitulate CEBPB-driven phenotypes, in response to autophagy inhibition. Hemin, or a γ-Secretase inhibitor counteract these phenotypes in CRISPR-C19MC-engineered cells. Therefore, a CEBPB and C19MC-driven M on -gene-signature regulates the podosomal belt, lipid droplet, HBV, and DEPTOR mRNA dynamics to genetically link obesity, and neurodegeneration at the cellular level.

Background Macrophages adopt activation states along a spectrum from pro- to anti-inflammatory, enabling appropriate responses to pathogens and environmental cues. Dysregulated inflammatory macrophage activation contributes to diseases including sepsis, rheumatoid arthritis, cancer, and atherosclerosis. Epigenetic processes such as DNA methylation and histone modification prime macrophages for activation, and several histone modifying enzymes (HMEs) have been implicated in this regulation. Objective To systematically identify histone modifying enzymes that regulate inflammatory macrophage activation. Methods We performed a CRISPR knockout screen with single-cell RNA-seq readout (CROP-seq) targeting 92 macrophage-expressed HMEs in immortalized LPS-activated mouse bone marrow-derived macrophages (BMDMs). The resulting single-cell transcriptomes were analyzed to identify significant perturbations. Kdm5c was selected for experimental validation in mouse BMDMs, and its expression pattern was compared with macrophage subsets from human atherosclerotic plaques using scRNA-seq data. Results The CROP-seq screen identified Prmt6, Carm1, Kat2b , and Kdm5c as top regulators of inflammatory macrophage activation. Validation in a KO cell line revealed loss of Kdm5c suppressed inflammatory tone at baseline but led to an exaggerated transcriptional response to LPS stimulation, indicating a role for Kdm5c in balancing tonic and inducible activation. A weighted gene module derived from Kdm5c -deficient macrophages was enriched in inflammatory macrophages in human atherosclerotic plaques. Conclusion Our findings demonstrate the value of CROP-seq screening to dissect the epigenetic control of macrophage activation. We also identify Kdm5c-mediated histone demethylation as a key mechanism modulating inflammatory macrophage activation.

NUT carcinoma (NC) is an aggressive malignancy driven by NUTM1 gene rearrangements with limited therapeutic options. Here, we show that direct suppression of NUTM1 using CRISPR/Cas9 induces squamous-like differentiation and upregulates TROP2 expression in NC cells. Building on this finding, we developed a TROP2–interferon beta (IFN-β) mutein immunocytokine that selectively targets TROP2-expressing tumors. Combined NUTM1 suppression and TROP2-targeted immunotherapy synergistically enhanced cytotoxic and immune-mediated responses in vitro . Transcriptomic and spatial analyses of NC patient tumors revealed that differentiation status correlates with TROP2 expression, upregulated immune pathways, and favorable clinical outcomes. Our results suggest that overcoming differentiation blockade not only alters tumor phenotype but also creates a more immune-permissive microenvironment. These findings highlight the therapeutic potential of sequential tumor reprogramming followed by targeted immunotherapy in treating NC and propose a broader strategy for overcoming differentiation blockade in fusion-driven cancers.

Acute myeloid leukemia (AML) is a heterogeneous disease that arises from dysregulated myeloid proliferation. We performed a high-throughput CRISPR interference (CRISPRi) screen in the THP-1 monocytic cancer cell line to identify long noncoding RNAs (lncRNAs) that play a role in contributing to cell proliferation. Our screen identified INSTAR (Intergenic Nuclear Suppressor lncRNA Targeting Adjacent Regulator SFMBT2 ) as a top candidate. RNA-seq on INSTAR deficient THP-1 cells revealed transcriptional changes in genes involved in cell proliferation as well as other cellular processes. Loss of INSTAR selectively reduced expression of its neighboring gene, SFMBT2 . Functional assays confirmed that both genes suppress cell growth, revealing a cis-regulatory mechanism in which INSTAR regulates SFMBT2 expression to control monocyte proliferation. Here, we leverage high-throughput screening to rapidly pinpoint functional lncRNAs providing novel insights into a key regulatory locus consisting of INSTAR and SFMBT2 which could be critical for better understanding dysregulation contributing to acute myeloid leukemia.

ABSTRACT CRISPR/Cas9-based mosaic analysis is a powerful tool for in vivo genetics but is limited by cytotoxicity and mutagenesis associated with DNA double-strand breaks (DSBs). Here, we establish Cas9-derived nickases as safer and more reliable alternatives for inducing mitotic recombination in Drosophila . We demonstrate that single-strand nicks are sufficient to generate mosaic clones and systematically dissect the parameters governing this process. We find that clone frequency can be controlled by the gRNA nicking pattern, with two distant nicks on the same DNA strand synergistically enhancing recombination by over nine-fold compared to a single nick. Based on these findings, we propose a mechanistic model for nick-induced crossover and provide a versatile toolkit for generating tissue-specific nickases. This work establishes nickase-based MAGIC as a superior method for high-fidelity clonal analysis, enabling more precise investigation of gene function in development and disease. SIGNIFICANCE STATEMENT The CRISPR/Cas9-based mosaic technique, MAGIC, is a versatile tool for in vivo biological investigations. However, its reliance on DNA double-strand breaks (DSBs) can cause significant, unintended cell damage. Here we establish that Cas9-derived nickases, which create gentler single-strand nicks, are a superior alternative. We show that nickases safely induce genetic mosaics in Drosophila by avoiding this cellular toxicity. By systematically dissecting the process, we discovered principles of gRNA design that allow clone frequencies to be ‘tuned’ for different experimental needs. This work provides a new mechanistic model for nick-induced genetic exchange, a high-fidelity “nickase-MAGIC” method, and a versatile toolkit for precision clonal analysis.

ABSTRACT The high mortality associated with tuberculosis (TB), alongside the lack of efficient therapeutics against emerging multidrug-resistant Mycobacterium tuberculosis ( Mtb ) strains, emphasizes the need to identify novel antitubercular targets. Mycobacterial peptidoglycan, displaying characteristic modifications comprising the amidation of D- iso -glutamate (D- i Glu) and the N -glycolylation of muramic acid, is a promising therapeutic target. The genes encoding the enzymes mediating these PG modifications ( murT / gatD and namH ) were silenced in Mtb using CRISPR interference (CRISPRi) to investigate their impact on β-lactam susceptibility and host immune responses. First, qRT-PCR confirmed successful target mRNA knockdown, with variable repression efficiency based on the selected sgRNA, PAM strength, and target site. Phenotypic characterization through spotting dilution and growth curve assays corroborated the essentiality of D- i Glu amidation for mycobacterial survival, in contrast to the N -glycolylation of muramic acid. Moreover, susceptibility assays showed that both PG modifications contribute to β-lactam resistance, with sgRNA2-mediated murT knockdown substantially increasing β-lactam and isoniazid susceptibility. Furthermore, checkerboard assays showed reductions in the minimum fractional inhibitory concentration index (FICI min ) value for AMX/MEM+CLA and EMB combinations following the depletion of both PG modifications, with significant differences observed upon namH knockdown. Additionally, D- i Glu amidation was uncovered as a determinant of Mtb survival within THP-1-derived macrophages at 6 days post-infection. Infection of THP-1-derived macrophages with MurT/GatD-depleted Mtb upregulated IL-1β and downregulated IL-10, whereas NamH depletion caused upregulation of both IL-1β and IL-10. Altogether, our findings unveiled the potential of targeting these PG modifications for the development of innovative therapeutic regimens against TB.

Abstract Genome editing with CRISPR-Cas9 offers a powerful approach for enhancing enzyme production in microorganisms. This study aimed to genetically engineer the lacZ gene in Escherichia coli using CRISPR-Cas9 to evaluate its impact on asparaginase production during submerged fermentation with rice bran serving as a glucose source. Both edited and wild-type E. coli strains were cultured at optimal conditions to produce and characterize asparaginase. The edited E. coli formed distinct colonies, displaying a blue phenotype when exposed to Cas9 without sgRNA or arabinose, yielding a total of 96 colonies. No colonies were observed when Cas9 and sgRNA were present without arabinose, while the addition of Cas9 and arabinose without sgRNA resulted in 309 blue colonies. With Cas9, sgRNA, and arabinose present, repair activation produced 114 distinct white colonies. The editing of the lacZ gene was validated through multiplex PCR and gel electrophoresis, with bands at 650 bp indicated lacZ gene editing, while bands at 1,100 bp indicated the wild-type. Asparaginase production was assessed using plate method assay, submerged fermentation using rice bran as a glucose source, and subsequent purification via ammonium sulfate precipitation and ion-exchange chromatography. Ion-exchange chromatography revealed enhanced purity and activity in the edited strain, with peak activity observed at an elution of 80 mL. The CRISPR-Cas9 edited strain exhibiting significantly higher enzyme activity (1.2 ± 0.002 U/ mL) compared to the wild-type (0.8 ± 0.005 U/mL). Both strains demonstrated maximum asparaginase activity at 40 o C and pH 7. This study concludes that CRISPR-Cas9 meditated lacZ gene editing in E. coli improves its ability to utilize rice bran as a substrate, significantly enhancing asparaginase production. These findings highlight the potential of genetic engineering and agricultural by-products for sustainable enzyme production.