Cell Biology

Diabetes lacks concrete curative strategies due to diverse aetiologies and, therefore, represents the perfect candidate for cell replacement therapy, since it is caused by either an absolute (type 1 diabetes) or relative (type 2 diabetes) defect in the insulin-producing beta cells of the pancreas. Pancreatic alpha cells are a promising source for transdifferentiation into insulin-producing cells as they share a common developmental origin with beta cells and exhibit a certain degree of cellular plasticity. Furthermore, impairment of glucagon signaling in diabetes leads to a marked increase in alpha cell mass, raising the possibility that such alpha cell hyperplasia provides an increased supply of alpha cells for their transdifferentiation into new beta cells.

In this protocol, we used the modular epigenetic CRISPR/dCas9 toolbox for targeted DNA methylation (EpiCRISPR) and silencing of the Arx gene (Aristaless Related Homeobox, Arx), which is essential for the maintenance of alpha cell identity. Methylation-based silencing of Arx initiates the reprogramming of pancreatic alpha cells into insulin-producing cells. As a key novelty, this protocol provides a direct route for epigenetically induced transdifferentiation of mouse pancreatic alpha TC1-6 cells into insulin-producing cells and thereby confirms a proof of concept of reversible cellular epigenetic reprogramming in vitro. In addition, this streamlined workflow addresses the inherent challenges of transfecting clustered alpha TC1-6 cells by optimizing their dissociation into single-cell suspensions, thereby improving uptake and reproducibility.

In summary, this approach for cell transdifferentiation involves precise epigenetic editing of a lineage-specific marker gene, thereby enabling direct lineage conversion in a safe and versatile strategy to generate insulin-producing cells by epigenetic reprogramming. In contrast to approaches that rely on viral vectors or permanent genome editing, this method reduces the risk of off-target effects and immunogenic responses while ensuring reproducibility. The combination of efficiency and precision makes it a valuable tool to advance regenerative approaches for diabetes therapy and to explore the epigenetic regulation of cell identity.

Adipocytes are endocrine cells that function as the main energy storage in our body. They are commonly used in clinical procedures, including their removal via liposuction and transplantation in plastic surgery. Building on this, adipocytes can be used for ex vivo cellular manipulations, enabling therapeutic modifications that can provide beneficial clinical outcomes after transplantation. Here, we provide a detailed protocol on how to modify adipocytes and adipose organoids using CRISPR activation (CRISPRa), a technology termed adipose manipulation transplantation (AMT).

Targeted genome editing of human pluripotent stem cells (hPSCs) is critical for basic and translational research and can be achieved with site-specific endonucleases. Cpf1 (CRISPR from Prevotella and Francisella) is a programmable DNA endonuclease with AT-rich PAM sequences. In this protocol, we describe procedures for using a single vector system to deliver Cpf1 and CRISPR RNA (crRNA) for genome editing in hPSCs. This protocol enables indel formation and homologous recombination–mediated precise editing at multiple loci. With the delivery of Cpf1 and a single U6 promoter-driven guide RNA array composed of an AAVS1-targeting and a MAFB-targeting crRNA array, efficient multiplex genome editing at the AAVS1 (knockin) and MAFB (knockout) loci in hPSCs could be achieved in a single experiment. The edited hPSCs expressed pluripotency markers and could differentiate into neurons in vitro. This system also generated INS reporter hPSCs with a 6 kb cassette knockin at the INS locus. The INS reporter cells can differentiate into β-cells that express tdTomato and luciferase, permitting fluorescence-activated cell sorting of hPSC-β-cells. By targeted screening of potential off-target sequences that are most homologous to crRNA sequences, no off-target mutations were detected in any of the tested sequences. This work provides an efficient and flexible system for precise genome editing in mammalian cells including hPSCs with the benefits of less off-target effects.

The adoptive transfer of autologous, long-lived, gene-repaired T cells is a promising way to treat inherited T-cell immunodeficiencies. However, adoptive T-cell therapies require a large number of T cells to be manipulated and infused back into the patient. This poses a challenge in primary immunodeficiencies that manifest early in childhood and where only small volumes of blood samples may be available. Our protocol describes the ex vivo expansion of potentially long-lived human T memory stem cells (T_SCM), starting from a limited number of peripheral blood mononuclear cells (PBMCs). Using the perforin gene as an example, we provide detailed instructions for precise gene repair of human T cells and the expansion of T_SCM. The efficiency of precise gene repair can be increased by suppressing unintended non-homologous end-joining (NHEJ) events. Our protocol yields edited T-cell populations that are ready for phenotyping, genome-wide off-target analysis, and functional characterization.

CRISPR/Cas9 genome editing is a widely used tool for creating genetic knock-ins, which allow for endogenous tagging of genes. This is in contrast with random insertion using viral vectors, where expression of the inserted transgene changes the total copy number of a gene in a cell and does not reflect the endogenous chromatin environment or any trans-acting regulation experienced at a locus. There are very few protocols for endogenous fluorescent tagging in macrophages. Here, we describe a protocol to design and test CRISPR guide RNAs and donor plasmids, to transfect them into RAW 264.7 mouse macrophage-like cells using the Neon transfection system and to grow up clonal populations of cells containing the endogenous knock-in at various loci. We have used this protocol to create endogenous fluorescent knock-ins in at least six loci, including both endogenously tagging genes and inserting transgenes in the Rosa26 and Tigre safe harbor loci. This protocol uses circular plasmid DNA as the donor template and delivers the sgRNA and Cas9 as an all-in-one expression plasmid. We designed this protocol for fluorescent protein knock-ins; it is best used when positive clones can be identified by fluorescence. However, it may be possible to adapt the protocol for non-fluorescent knock-ins. This protocol allows for the fairly straightforward creation of clonal populations of macrophages with tags at the endogenous loci of genes. We also describe how to set up imaging experiments in 24-well plates to track fluorescence in the edited cells over time.

Key features

• CRISPR knock-in of fluorescent proteins in RAW 264.7 mouse macrophages at diverse genomic loci.

• This protocol is optimized for the use of the Neon transfection system.

• Includes instructions for growing up edited clonal populations from single cells with one single-cell sorting step and efficient growth in conditioned media after cell sorting.

• Designed for knocking in fluorescent proteins and screening transfected cells byFACS, but modification for non-fluorescent knock-ins may be possible.

Graphical overview

An efficient and precise genome-editing approach is in high demand in any molecular biology or cell biology laboratory worldwide. However, despite a recent rapid progress in the toolbox tailored for precise genome-editing, including the base editors and prime editors, there is still a need for a cost-effective knock-in (KI) approach amenable for long donor DNA cargos with high efficiency. By harnessing the high-efficient double-strand break (DSB) repair pathway of microhomology-mediated end joining, we previously showed that a specially designed 3′-overhang double-strand DNA (odsDNA) donor harboring 50-nt homology arm (HA) allows high-efficient exogenous DNA KI when combined with CRISPR-Cas9 technology. The lengths of the 3′-overhangs of odsDNA donors could be manipulated by the five consecutive phosphorothioate (PT) modifications. In this protocol, we detail the stepwise procedures to conduct the LOCK (Long dsDNA with 3′-Overhangs mediated CRISPR Knock-in) method for gene-sized (~1–3 kb) KI in mammalian cells.

Graphical overview

Improvement of large DNA fragment knock-in rates by attaching odsDNA donors to Cas9-PCV2 fusion protein

Synapses are specialized structures that enable neuronal communication, which is essential for brain function and development. Alterations in synaptic proteins have been linked to various neurological and neuropsychiatric disorders. Therefore, manipulating synaptic proteins in vivo can provide insight into the molecular mechanisms underlying these disorders and aid in developing new therapeutic strategies. Previous methods such as constitutive knock-out animals are limited by developmental compensation and off-target effects. The current approach outlines procedures for age-dependent molecular manipulations in mice using helper-dependent adenovirus viral vectors (HdAd) at distinct developmental time points. Using stereotactic injection of HdAds in both newborn and juvenile mice, we demonstrate the versatility of this method to express Cre recombinase in globular bushy cells of juvenile Rac1^fl/fl mice to ablate presynaptic Rac1 and study its role in synaptic transmission. Separately, we overexpress Ca_V2 α1 subunits at two distinct developmental time points to elucidate the mechanisms that determine presynaptic Ca_V2 channel abundance and preference. This method presents a reliable, cost-effective, and minimally invasive approach for controlling gene expression in specific regions of the mouse brain and will be a powerful tool to decipher brain function in health and disease.

Key features

• Virus-mediated genetic perturbation in neonatal and young adult mice.

• Stereotaxic injection allows targeting of brain structures at different developmental stages to study the impact of genetic perturbation throughout the development.

Development of the hybridoma technology by Köhler and Milstein (1975) has revolutionized the immunological field by enabling routine use of monoclonal antibodies (mAbs) in research and development efforts, resulting in their successful application in the clinic today. While recombinant good manufacturing practices production technologies are required to produce clinical grade mAbs, academic laboratories and biotechnology companies still rely on the original hybridoma lines to stably and effortlessly produce high antibody yields at a modest price. In our own work, we were confronted with a major issue when using hybridoma-derived mAbs: there was no control over the antibody format that was produced, a flexibility that recombinant production does allow. We set out to remove this hurdle by genetically engineering antibodies directly in the immunoglobulin (Ig) locus of hybridoma cells. We used clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) and homology-directed repair (HDR) to modify antibody’s format [mAb or antigen-binding fragment (Fab’)] and isotype. This protocol describes a straightforward approach, with little hands-on time, leading to stable cell lines secreting high levels of engineered antibodies. Parental hybridoma cells are maintained in culture, transfected with a guide RNA (gRNA) targeting the site of interest in the Ig locus and an HDR template to knock in the desired insert and an antibiotic resistance gene. By applying antibiotic pressure, resistant clones are expanded and characterized at the genetic and protein level for their ability to produce modified mAbs instead of the parental protein. Finally, the modified antibody is characterized in functional assays. To demonstrate the versatility of our strategy, we illustrate this protocol with examples where we have (i) exchanged the constant heavy region of the antibody, creating chimeric mAb of a novel isotype, (ii) truncated the antibody to create an antigenic peptide-fused Fab’ fragment to produce a dendritic cell–targeted vaccine, and (iii) modified both the constant heavy (CH)1 domain of the heavy chain (HC) and the constant kappa (Cκ) light chain (LC) to introduce site-selective modification tags for further derivatization of the purified protein. Only standard laboratory equipment is required, which facilitates its application across various labs. We hope that this protocol will further disseminate our technology and help other researchers.

Graphical abstract

Chemical modifications on RNA play important roles in regulating its fate and various biological activities. However, the impact of RNA modifications varies depending on their locations on different transcripts and cells/tissues contexts; available tools to dissect context-specific RNA modifications are still limited. Herein, we report the detailed protocol for using a chemically inducible and reversible platform to achieve site-specific editing of the chosen RNA modification in a temporally controlled manner by integrating the clustered regularly interspaced short palindromic repeats (CRISPR) technology and the abscisic acid (ABA)-based chemically induced proximity (CIP) system. The procedures were demonstrated using the example of inducible and reversible N⁶-methyladenosine (m⁶A) editing and the evaluation of its impact on RNA properties with ABA addition and reversal with the control of ABA or light.

Loss-of-function (LoF) variants in the low-density lipoprotein receptor–related protein 10 gene (LRP10) have been recently implicated in the development of neurodegenerative diseases, including Parkinson's disease (PD), PD dementia (PDD), and dementia with Lewy bodies (DLB). However, despite the genetic evidence, little is known about the LRP10 protein function in health and disease. Here, we describe a detailed protocol to efficiently generate a LRP10 LoF model in two independent LRP10-expressing cell lines, HuTu-80 and HEK 293T, using the CRISPR/Cas9 genome-editing tool. Our method efficiently generates bi-allelic LRP10 knockout (KO), which can be further utilized to elucidate the physiological LRP10 protein function and to model some aspects of neurodegenerative disorders.

Graphical abstract:

CRISPR/Cas9 workflow for the generation of the LRP10 KO. (1) Designed single guide RNA (sgRNA) is cloned into CRISPR/Cas9 px458 plasmid. (2) Cells are transfected with the CRISPR/Cas9 plasmid containing sgRNA. (3) Two days post transfection, cells are dissociated and sorted as single cells by fluorescence-activated cell sorting (FACS). (4) After several weeks, expanded clonal lines are (5) verified with Sanger sequencing for the presence of INDELs (insertions or deletions), RT-qPCR for the amounts of LRP10 mRNA transcript, and Western blotting for the analysis of the LRP10 protein levels.