Cell Biology


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0 Q&A 774 Views Oct 20, 2023

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

0 Q&A 448 Views Aug 20, 2023

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 Rac1fl/fl mice to ablate presynaptic Rac1 and study its role in synaptic transmission. Separately, we overexpress CaV2 α1 subunits at two distinct developmental time points to elucidate the mechanisms that determine presynaptic CaV2 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.

0 Q&A 730 Views Feb 20, 2023

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

0 Q&A 491 Views Feb 5, 2023

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.

0 Q&A 1818 Views Oct 5, 2022

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.

0 Q&A 1931 Views May 20, 2022

Although CRISPR-Cas9 genome editing can be performed directly in single-cell mouse zygotes, the targeting efficiency for more complex modifications such as the insertion of two loxP sites, multiple mutations in cis, or the precise insertion or deletion of longer DNA sequences often remains low (Cohen, 2016). Thus, targeting and validation of correct genomic modification in murine embryonic stem cells (ESCs) with subsequent injection into early-stage mouse embryos may still be preferable, allowing for large-scale screening in vitro before transfer of thoroughly characterized and genetically defined ESC clones into the germline. This procedure can result in a reduction of animal numbers with cost effectiveness and compliance with the 3R principle of animal welfare regulations. Here, we demonstrate that after transfection of homology templates and PX458 CRISPR-Cas9 plasmids, EGFP-positive ESCs can be sorted with a flow cytometer for the enrichment of CRISPR-Cas9-expressing cells. Cell sorting obviates antibiotic selection and therefore allows for more gentle culture conditions and faster outgrowth of ESC clones, which are then screened by qPCR for correct genomic modifications. qPCR screening is more convenient and less time-consuming compared to analyzing PCR samples on agarose gels. Positive ESC clones are validated by PCR analysis and sequencing and can serve for injection into early-stage mouse embryos for the generation of chimeric mice with germline transmission. Therefore, we describe here a simple and straightforward protocol for CRISPR-Cas9-directed gene targeting in ESCs.

Graphical abstract:

0 Q&A 3285 Views May 20, 2022

Genome editing by the delivery of pre-assembled Cas9 ribonucleoproteins (Cas9 RNP) is an increasingly popular approach for cell types that are difficult to manipulate genetically by the conventional plasmid and viral methods. Cas9 RNP editing is robust, precise, capable of multiplexing, and free of genetic materials. Its transient presence in cells limits residual editing activity. This protocol describes the preparation of recombinant Streptococcus pyogenes Cas9 (SpCas9) protein by heterologous expression and purification from Escherichia coli, and the synthesis of CRISPR guide RNA by in vitro transcription and PAGE purification. SpCas9 is the first CRISPR Cas9 discovered (Jinek et al., 2012) and is also one of the most characterized Cas enzymes for genome editing applications. Using this formulation of Cas9 RNP, we have demonstrated highly efficient genome editing in primary human T and natural killer (NK) cells by electroporation, and in fungi and plants by polyethylene glycol-mediated transformation. Our protocol of Cas9 RNP preparation is consistent and straightforward to adopt for genome editing in other cell types and organisms.

Graphical abstract:

0 Q&A 2325 Views May 20, 2022

Subcellular localization dynamics of proteins involved in signal transduction processes is crucial in determining the signaling outcome. However, there is very limited information about the localization of endogenous signaling proteins in living cells. For example, biochemical mechanisms underlying the signaling pathway from epidermal growth factor (EGF) receptor (EGFR) to RAS-RAF and ERK1/2/MAPK are well understood, whereas the operational domains of this pathway in the cell remain poorly characterized. Tagging of endogenous components of signaling pathways with fluorescent proteins allows more accurate characterization of their intracellular dynamics at their native expression levels controlled by endogenous regulatory mechanisms, thus avoiding possible tainting effects of overexpression and mistargeting. In this study, we describe methodological approaches to label components of the EGFR-RAS-MAPK pathway, such as Grb2, KRAS, and NRAS, with the fluorescent protein mNeonGreen (mNG) using CRISPR/Cas9 gene-editing, as well as generation of homozygous single-cell clones of the edited target protein.

0 Q&A 2276 Views Mar 5, 2022

Directed evolution is a powerful approach to obtain genetically-encoded sought-for traits. Compared to the prolonged adaptation regimes to mutations occurring under natural selection, directed evolution unlocks rapid screening and selection of mutants with improved traits from vast mutated sequence spaces. Many systems have been developed to search variant landscapes based on ex vivo or in vivo mutagenesis, to identify and select new-to-nature and optimized properties in biomolecules. Yet, the majority of such systems rely on tedious iterations of library preparation, propagation, and selection steps. Furthermore, among the relatively few in vivo directed evolution systems developed to mitigate handling of repetitive ex vivo steps, directed evolution of DNA is the standard approach. Here, we present the protocol for designing the transfer of genetic information from evolving RNA donors to DNA in baker’s yeast, using CRISPR- and RNA-assisted in vivo directed evolution (CRAIDE). We use mutant T7 RNA polymerase to introduce mutations in RNA donors, while incorporation into DNA is directed by CRISPR/Cas9. As such, CRAIDE offers an opportunity to study fundamental questions, such as RNA’s contribution to the evolution of DNA-based life on Earth.

Graphic abstract:

CRISPR- and RNA-assisted in vivo directed evolution (CRAIDE).

0 Q&A 6607 Views Mar 5, 2022

The CRISPR/Cas9 technology has transformed our ability to edit eukaryotic genomes. Despite this breakthrough, it remains challenging to precisely knock-in large DNA sequences, such as those encoding a fluorescent protein, for labeling or modifying a target protein in post-mitotic cells. Previous efforts focusing on sequence insertion to the protein coding sequence often suffer from insertions/deletions (INDELs) resulting from the efficient non-homologous end joining pathway (NHEJ). To overcome this limitation, we have developed CRISPR-mediated insertion of exon (CRISPIE). CRISPIE circumvents INDELs and other editing errors by inserting a designer exon flanked by adjacent intron sequences into an appropriate intronic location of the targeted gene. Because INDELs at the insertion junction can be spliced out, “CRISPIEd” genes produce precisely edited mRNA transcripts that are virtually error-free. In part due to the elimination of INDELs, high-efficiency labeling can be achieved in vivo. CRISPIE is compatible with both N- and C-terminal labels, and with all common transfection methods. Importantly, CRISPIE allows for later removal of the protein modification by including exogenous single-guide RNA (sgRNA) sites in the intronic region of the donor module. This protocol provides the detailed CRISPIE methodology, using endogenous labeling of β-actin in human U-2 OS cells with enhanced green fluorescent protein (EGFP) as an example. When combined with the appropriate gene delivery methods, the same methodology can be applied to label post-mitotic neurons in culture and in vivo. This methodology can also be readily adapted for use in other gene editing contexts.

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