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Tailoring microbial fitness through computational steering and CRISPRi-driven robustness regulation.
The widespread application of genetically modified microorganisms (GMMs) across diverse sectors underscores the pressing need for robust strategies to mitigate the risks associated with their potential uncontrolled escape. This study merges computational modeling with CRISPR interference (CRISPRi) to refine GMM metabolic robustness. Utilizing ensemble modeling, we achieved high-throughput in silico screening for enzymatic targets susceptible to expression alterations. Translating these insights, we developed functional CRISPRi, boosting fitness control via multiplexed gene knockdown. Our method, enhanced by an insulator-improved gRNA structure and an off-switch circuit controlling a compact Cas12m, resulted in rationally engineered strains with escape frequencies below National Institutes of Health standards. The effectiveness of this approach was confirmed under various conditions, showcasing its ability for secure GMM management. This research underscores the resilience of microbial metabolism, strategically modifying key nodes to halt growth without provoking significant resistance, thereby enabling more reliable and precise GMM control. A record of this paper's transparent peer review process is included in the supplemental information.
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Use of CRISPR/Cas9 for Targeted Mutagenesis in Sorghum.
Sorghum (Sorghum bicolor) fulfills the demand for bioenergy resources and also provides substantial diet calories to the world's population. Therefore, many biological studies use sorghum as a research model for improvement of the domesticated food and bioenergy crops. Furthermore, leveraging genome editing systems in a plethora of grass plant species has been extensively studied with no exception in sorghum. However, a protocol that details the genome editing strategies using CRISPR/Cas9 and that combines an efficient tissue culture and transformation platform in sorghum based on Agrobacterium-mediated DNA transfer has yet to be reported. This protocol outlines the steps and workflow from design of sorghum CRISPR target sites using BTx623 as a reference genome, construction of sorghum CRISPR/Cas9 plasmids, tissue culture, to Agrobacterium-mediated transformation followed by genotyping of CRISPR/Cas9 induced mutants. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Construction of CRISPR/Cas9 expression vector to analysis of CRISPR-edited plants Basic Protocol 2: Stable transformation of sorghum Support Protocol: Management of sorghum plants in a greenhouse.
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Quantitative Analysis of CRISPR Encoding Plasmid DNA-Nanocarrier Interactions by Gel Electrophoresis and Densitometry.
Nanoparticle-based carriers are key systems for the transport and delivery of nucleic acids in gene therapy, RNA therapeutics, and genome editing applications. The development of efficient delivery systems for CRISPR-Cas technologies requires reliable methods to evaluate nanocarrier-nucleic acid interactions, binding efficiency, and loading capacity. Agarose gel electrophoresis is commonly used to study these interactions through gel retardation assays, but results are often interpreted qualitatively based on visual inspection of DNA band migration. In this chapter, we present a quantitative approach that combines agarose gel electrophoresis with densitometric analysis of plasmid DNA (pDNA) bands to evaluate the interaction between pDNA and inorganic nanocarriers. The method relies on quantifying the fraction of free pDNA that migrates through the gel, while nanocarrier-pDNA systems remain near the loading wells. Band intensities are analyzed using ImageJ software to determine the amount of free pDNA and to estimate binding efficiency through depletion analysis. This protocol provides a simple and accessible strategy to quantify the loading of pDNA encoding CRISPR components, guide RNA constructs, or other nucleic acid cargos, facilitating the development and optimization of nanoparticle-based delivery systems for genome editing applications.
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CRISPR/Cas9 in zebrafish: an efficient combination for human genetic diseases modeling.
The next-generation sequencing identifies a growing number of candidate genes associated with human genetic diseases, which inevitably requires efficient methods to validate the causal links between genotype and phenotype. Recently, zebrafish, with sufficiently high-throughput capabilities, has become a favored option to study human pathogenesis. In addition, CRISPR/Cas9-based approaches have radically reduced the efforts to introduce targeted genome engineering in various organisms. Here, we systemically review the basic considerations in the design of gene editing in zebrafish with CRISPR/Cas9, and explore the potential of the combination of these two to support efficient functional analysis of human genetic variants.
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Harnessing CRISPR/Cas9 technology in cardiovascular disease.
The CRISPR/Cas9 system is a precisely targeted bacterial defense system, used to control invading viruses. This technology has many potential applications including genetic changes in somatic and germ cells and the creation of knockout animals. Compared to other genome editing techniques such as zinc-finger nucleases and transcription activator-like effector nucleases (TALENS), the CRISPR/Cas9 system is much easier and more efficient. Most importantly, the multifunctional capacity of this technology allows simultaneous editing of several genes. The CRISPR/Cas9 system also potentially has the ability to prevent and treat human diseases. The present article addresses some key points related to the use of the CRISPR/Cas9 system as a powerful tool in cardiovascular research and as a new strategy for the treatment of cardiovascular disease (CVD).
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BAR-CAT: Targeted Recovery of Synthetic Genes via Barcode-Directed CRISPR-dCas9 Enrichment.
Modern gene synthesis platforms enable investigations of protein function and genome biology at an unprecedented scale. Yet, the proportion of error-free constructs in diverse gene libraries decreases with length due to the propagation of oligo synthesis errors. To rescue these error-free constructs, we developed Barcode-Assisted Retrieval CRISPR-Activated Targeting (BAR-CAT), an in vitro method that uses multiplexed dCas9-single-guide RNA (sgRNA) complexes to extract barcodes corresponding to error-free constructs. After a 15-min incubation and wash regimen, three low-bundance targets in a 300,000-member test library were enriched 600-fold, greatly reducing downstream requirements. When applied to a 384-gene DropSynth gene library, BAR-CAT enriched 12 targets up to 122-fold and revealed practical limits imposed by sgRNA competition and library complexity, which now guide ongoing protocol scaling. By eliminating laborious clone-by-clone validation and working directly on plasmid libraries, BAR-CAT provides a platform for recovering perfect synthetic genes, subsetting large libraries, and ultimately lowering the cost of functional genomics at scale.
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CRISPR-Hybrid: Intracellular Selection of CRISPR-Associated Aptamers in Bacteria.
CRISPR technologies have evolved from nuclease-based genome editing to programmable systems for transcriptional and epigenetic regulations. Emerging CRISPR systems expand editing versatility by incorporating CRISPR-associated aptamers (CAPs) into single-guide RNAs (sgRNAs), enabling recruitment of RNA-binding proteins (RBPs) fused to diverse effectors. However, the limited availability of orthogonal aptamer-RBP pairs has hindered broad application, as conventional SELEX-based aptamer discovery is time-intensive and often fails to yield aptamers functional in cells. We developed the CRISPR-Hybrid platform, an intracellular selection method that directly evolves CAPs within bacterial cells. This system links aptamer-RBP interactions to a fluorescent reporter readout, allowing fluorescence-activated cell sorting (FACS) to enrich functional variants from libraries exceeding 10⁷ sequences. Each selection round can be completed in 2 days, enabling rapid enrichment of aptamers that retain activity in both bacterial and mammalian contexts. This protocol details construction of randomized DNA libraries, preparation of host cells, execution of intracellular selection and FACS enrichment, and recovery of aptamer sequences for downstream analysis. By providing a fast, in-cell, and physiologically relevant approach to aptamer discovery, CRISPR-Hybrid expands the repertoire of CAPs available for modular and multiplexed CRISPR editing.
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Initial Characterization of 12 New Subtypes and Variants of Type V CRISPR Systems.
Type V CRISPR systems are highly diverse in sequence, mechanism, and function. Although recent efforts have greatly expanded our understanding of their evolution, the diversity of type V systems remains to be completely explored, and many clades have not been experimentally characterized. In this work, we mined metagenomic databases to identify three new subtypes and nine new variants of Cas12, the effector of Type V systems, and provide experimental and computational characterization of their Protospacer-Adjacent Motif (PAM), interference activity, loci architecture, and tracrRNA dependence. Half of the new Cas12s are found in phages or prophages. New subtypes Cas12o and Cas12p lack the canonical RuvC catalytic residues, suggesting they interfere with the target without cleavage, possibly by blocking transcription or replication. One variant, Cas12f10, displays substantial activity on PAM-less targets. Our work expands the diversity of the functionally characterized Cas12 effectors and provides some promising candidates for genome engineering tools.
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AI-driven CRISPR screening: optimizing gene editing through automation and intelligent decision support.
BACKGROUND: CRISPR-based genetic screening has become a central methodology in functional genomics, enabling systematic interrogation of gene function, genetic interactions and context-dependent vulnerabilities at scale. However, the rapid expansion of screening modalities-including multi-condition designs, combinatorial perturbations, in vivo applications and single-cell readouts-has exposed fundamental limitations of heuristic-driven experimental design and post hoc statistical analysis. MAIN BODY: This Review synthesizes how artificial intelligence is reshaping CRISPR screening by introducing predictive, adaptive and system-level intelligence across the experimental lifecycle. We organize recent advances into two tightly coupled modules. First, machine learning and deep learning (ML/DL) methods optimize experimental design by learning context-dependent perturbation behavior, anticipating confounding effects and enabling iterative, information-efficient screening strategies. Second, large language model-agent (LLM-agent) systems complement these advances by externalizing scientific reasoning, integrating biological knowledge at scale and coordinating analysis and decision-making in human-in-the-loop workflows. CONCLUSIONS: Together, ML/DL and LLM-agent approaches reframe CRISPR screening from a static analytical pipeline into an intelligent experimental system, with important implications for robustness, scalability and biological discovery.
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Advanced genome-editing technologies enable rapid and large-scale generation of genetic variants for strain engineering and synthetic biology.
Targeted genome editing not only improves our understanding of fundamental rules in life sciences but also affords us versatile toolkits to improve industrially relevant phenotypes in various host cells. In this review, we summarize the recent endeavor to develop efficient genome-editing tools, and emphasize the utility of these tools to generate massive scale of genetic variants. We categorize these tools into traditional recombination-based tools, and more advanced CRISPR as well as RNA-based genome-editing tools. This diverse panel of sophisticated tools has been applied to accelerate strain engineering, upgrade biomanufacturing, and customize biosensing. In parallel with high-throughput phenotyping and AI-based optimization algorithms, we envision that genome-editing technologies will become a driving force to automate and streamline biological engineering, and empower us to address critical challenges in health, environment, energy, and sustainability.
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