Cell Biology

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

Cancer cells evade the immune system by downregulating antigen presentation. Although immune checkpoint inhibitors (ICI) and adoptive T-cell therapies revolutionized cancer treatment, their efficacy relies on the intrinsic immunogenicity of tumor cells and antigen presentation by dendritic cells. Here, we describe a protocol to directly reprogram murine and human cancer cells into tumor-antigen-presenting cells (tumor-APCs), using the type 1 conventional dendritic cell (cDC1) transcription factors PU.1, IRF8, and BATF3 delivered by a lentiviral vector. Tumor-APCs acquire a cDC1 cell-like phenotype, transcriptional and epigenetic programs, and function within nine days (Zimmermannova et al., 2023). Tumor-APCs express the hematopoietic marker CD45 and acquire the antigen presentation complexes MHC class I and II as well as co-stimulatory molecules required for antigen presentation to T cells, but do not express high levels of negative immune checkpoint regulators. Enriched tumor-APCs present antigens to Naïve CD8⁺ and CD4⁺ T cells, are targeted by activated cytotoxic T lymphocytes, and elicit anti-tumor responses in vivo. The tumor-APC reprogramming protocol described here provides a simple and robust method to revert tumor evasion mechanisms by increasing antigen presentation in cancer cells. This platform has the potential to prime antigen-specific T-cell expansion, which can be leveraged for developing new cancer vaccines, neoantigen discovery, and expansion of tumor-infiltrating lymphocytes.

Key features

• This protocol describes the generation of antigen-presenting cells from cancer cells by direct reprogramming using lineage-instructive transcription factors of conventional dendritic cells type I.

• Verification of reprogramming efficiency by flow cytometry and functional assessment of tumor-APCs by antigen presentation assays.

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.

Invariant natural killer T (iNKT) cells are a non-conventional T-cell population expressing a conserved semi-invariant T-cell receptor (TCR) that reacts to lipid antigens, such as α-galactosyl ceramide (α-GalCer), presented by the monomorphic molecule CD1d. iNKT cells play a central role in tumor immunosurveillance and represent a powerful tool for anti-cancer treatment, notably because they can be efficiently redirected against hematological or solid malignancies by engineering with tumor-specific chimeric antigen receptors (CARs) or TCRs. However, iNKT cells are rare and require specific ex vivo pre-selection and substantial in vitro expansion to be exploited for adoptive cell therapy (ACT). This protocol describes a robust method to obtain a large number of mouse iNKT cells that can be effectually engineered by retroviral (RV) transduction. A major advantage of this protocol is that it requires neither particular instrumentation nor a high number of mice. iNKT cells are enriched from the spleens of iVα14-Jα18 transgenic mice; the rapid purification protocol yields a highly enriched iNKT cell population that is activated by anti-CD3/CD28 beads, which is more reproducible and less time consuming than using bone marrow–derived dendritic cells loaded with α-GalCer, without risks of expanding contaminant T cells. Forty-eight hours after activation, iNKT cells are transduced with the selected RV by spin inoculation. This protocol allows to obtain, in 15 days, millions of ready-to-use, highly pure, and stably transduced iNKT cells that might be exploited for in vitro assays and ACT experiments in preclinical studies.

Three-dimensional bioprinting utilizes additive manufacturing processes that combine cells and a bioink to create living tissue models that mimic tissues found in vivo. Stem cells can regenerate and differentiate into specialized cell types, making them valuable for research concerning degenerative diseases and their potential treatments. 3D bioprinting stem cell–derived tissues have an advantage over other cell types because they can be expanded in large quantities and then differentiated to multiple cell types. Using patient-derived stem cells also enables a personalized medicine approach to the study of disease progression. In particular, mesenchymal stem cells (MSC) are an attractive cell type for bioprinting because they are easier to obtain from patients in comparison to pluripotent stem cells, and their robust characteristics make them desirable for bioprinting. Currently, both MSC bioprinting protocols and cell culturing protocols exist separately, but there is a lack of literature that combines the culturing of the cells with the bioprinting process. This protocol aims to bridge that gap by describing the bioprinting process in detail, starting with how to culture cells pre-printing, to 3D bioprinting the cells, and finally to the culturing process post-printing. Here, we outline the process of culturing MSCs to produce cells for 3D bioprinting. We also describe the process of preparing Axolotl Biosciences TissuePrint - High Viscosity (HV) and Low Viscosity (LV) bioink, the incorporation of MSCs to the bioink, setting up the BIO X and the Aspect RX1 bioprinters, and necessary computer-aided design (CAD) files. We also detail the differentiation of 2D and 3D cell cultures of MSC to dopaminergic neurons, including media preparation. We have also included the protocols for viability, immunocytochemistry, electrophysiology, and performing a dopamine enzyme-linked immunosorbent assay (ELISA), along with the statistical analysis.

Graphical overview

Genetic strategies such as gene disruption and fluorescent protein tagging largely contribute to understanding the molecular mechanisms of biological functions in bacteria. However, the methods for gene replacement remain underdeveloped for the filamentous bacteria Leptothrix cholodnii SP-6. Their cell chains are encased in sheath composed of entangled nanofibrils, which may prevent the conjugation for gene transfer. Here, we describe a protocol optimized for gene disruption through gene transfer mediated by conjugation with Escherichia coli S17-1 with details on cell ratio, sheath removal, and loci validation. The obtained deletion mutants for specific genes can be used to clarify the biological functions of the proteins encoded by the target genes.

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

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.