细胞生物学

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

Recombinant adeno-associated viruses (rAAVs) are valuable viral vectors for in vivo gene transfer, also having significant ex vivo therapeutic potential. Continued efforts have focused on various gene therapy applications, capsid engineering, and scalable manufacturing processes. Adherent cells are commonly used for virus production in most basic science laboratories because of their efficiency and cost. Although suspension cells are easier to handle and scale up compared to adherent cells, their use in virus production is hampered by poor transfection efficiency. In this protocol, we developed a simple scalable AAV production protocol using serum-free-media-adapted HEK293T suspension cells and VirusGEN transfection reagent. The established protocol allows AAV production from transfection to quality analysis of purified AAV within two weeks. Typical vector yields for the described suspension system followed by iodixanol purification range from a total of 1 × 10¹³ to 1.5 × 10¹³ vg (vector genome) using 90 mL of cell suspension vs. 1 × 10¹³ to 2 × 10¹³ vg using a regular adherent cell protocol (10 × 15 cm dishes).

Key features

• Adeno-associated virus (AAV) production using serum-free-media-adapted HEK293T suspension cells.

• Efficient transfection with VirusGEN.

• High AAV yield from small-volume cell culture.

Graphical overview

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

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.

Type 1 regulatory T (Tr1) cells are an immunoregulatory CD4⁺ Foxp3^- IL-10^high T cell subset with therapeutic potential for various inflammatory diseases. Retroviral (RV) transduction has been a valuable tool in defining the signaling pathways and transcription factors that regulate Tr1 differentiation and suppressive function. This protocol describes a method for RV transduction of naïve CD4⁺ T cells differentiating under Tr1 conditions, without the use of reagents such as polybrene or RetroNectin. A major advantage of this protocol over others is that it allows for the role of genes of interest on both differentiation and function of Tr1 cells to be interrogated. This is due to the high efficiency of RV transduction combined with the use of an IL10^GFP/Foxp3^RFP dual reporter mouse model, which enables successfully transduced Tr1 cells to be identified and sorted for functional assays. In addition, this protocol may be utilized for dual/multiple transduction approaches and transduction of other lymphocyte populations, such as CD8⁺ T cells.

Currently, there are several in vitro protocols that focus on directing human induced pluripotent stem cell (hiPSC) differentiation into either the cardiac or pulmonary lineage. However, these systemsprotocols are unable to recapitulate the critical exchange of signals and cells between the heart and lungs during early development. To address this gap, here we describe a protocol to co-differentiate cardiac and pulmonary progenitors within a single hiPSC culture by temporal specific modulation of Wnt and Nodal signaling. Subsequently, human cardio-pulmonary micro-tissues (μTs) can be generated by culturing the co-induced cardiac and pulmonary progenitors in 3D suspension culture. Anticipated results include expedited alveolarization in the presence of cardiac cells, and segregation of the cardiac and pulmonary μTs in the absence of exogenous Wnt signaling. This protocol can be used to model cardiac and pulmonary co-development, with potential applications in drug testing, and as a platform for expediting the maturation of pulmonary cells for lung tissue engineering.

There is an urgent need for the development of brain drug delivery carriers based on middle-sized or macromolecules, to which in vitro blood-brain barrier (BBB) models are expected to contribute significantly through evaluation of BBB permeability. As part of efforts to develop such models, we have been working on human conditionally immortalized cell-based multicellular spheroidal BBB models (hiMCS-BBB models), and we herein introduce the model development protocol. Briefly, astrocytes are first seeded in an ultra-low attachment 3D cell culture plate, to make the central core (Day 0). Next, pericytes are added over the core, to form an outer layer (Day 1). Then, brain microvascular endothelial cells are further added to each well, to create the outmost monolayer serving as the BBB (Day 2). Finally, the spheroids cultured for two days (on Day 4) can be used for assays of interest (e.g., antibody permeability assays). Neither special equipment nor techniques are required to produce hiMCS-BBB models. Therefore, the protocol presented here will not only facilitate the model sharing among the BBB community but also provide some technical clues contributing to the development of similar MCS-BBB models using other cell sources, such as primary or iPS-derived BBB cells.

Graphical abstract:

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:

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:

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.