Cancer Biology

CRISPR/Cas9 screening has revolutionized functional genomics in biomedical research and is a widely used approach for the identification of genetic dependencies in cancer cells. Here, we present an efficient and versatile protocol for the cloning of guide RNAs (gRNA) into lentiviral vectors, the production of lentiviral supernatants, and the transduction of target cells in a 96-well format. To assess the effect of gene knockouts on cellular fitness, we describe a competition-based cell proliferation assay using flow cytometry, enabling the screening of many genes at the same time in a fast and reproducible manner. This readout can be extended to any parameter that is accessible to flow-based measurements, such as protein expression and stability, differentiation, cell death, and others. In summary, this protocol allows to functionally assess the effect of a set of 50–300 gene knockouts on various cellular parameters within eight weeks.

Graphical abstract

N⁶-methyladenosine (m⁶A) is the most prevalent internal modification of eukaryotic messenger RNAs (mRNAs), affecting their fold, stability, degradation, and cellular interaction(s) and implicating them in processes such as splicing, translation, export, and decay. The m⁶A modification is also extensively present in non-coding RNAs, including microRNAs (miRNAs), ribosomal RNAs (rRNAs), and transfer RNAs (tRNAs). Common m6A methylation detection techniques play an important role in understanding the biological function and potential mechanism of m⁶A, mainly including the quantification and specific localization of m⁶A modification sites. Here, we describe in detail the dot blotting method for detecting m⁶A levels in RNA (mRNA as an example), including total RNA extraction, mRNA purification, dot blotting, and data analysis. This protocol can also be used to enrich specific RNAs (such as tRNA, rRNA, or miRNA) by isolation technology to detect the m⁶A level of single RNA species, so as to facilitate further studies of the role of m6A in biological processes.

The core planar cell polarity (PCP) protein Vang/Vangl, including Vangl1 and Vangl2 in vertebrates, is indispensable during development. Our previous studies showed that the activity of Vangl is tightly controlled by two important posttranslational modifications, ubiquitination and phosphorylation. Vangl is ubiquitinated through an endoplasmic reticulum-associated degradation (ERAD) pathway and is phosphorylated by casein kinase 1 (CK1) in response to Wnt. Here, we present step-by-step procedures to analyze Vangl ubiquitination and phosphorylation, including cell culture, transfection, sample preparation, and signal detection, as well as the use of newly available phospho-specific antibodies to detect Wnt-induced Vangl2 phosphorylation. The protocol described here can be applicable to the analysis of posttranslational modifications of other membrane proteins.

RNA binding proteins (RBPs) are critical regulators of cellular phenotypes, and dysregulated RBP expression is implicated in various diseases including cancer. A single RBP can bind to and regulate the expression of many RNA molecules via a variety of mechanisms, including translational suppression, prevention of RNA degradation, and alteration in subcellular localization. To elucidate the role of a specific RBP within a given cellular context, it is essential to first identify the group of RNA molecules to which it binds. This has traditionally been achieved using cross-linking-based assays in which cells are first exposed to agents that cross-link RBPs to nucleic acids and then lysed to extract and purify the RBP-nucleic acid complexes. The nucleic acids within the mixture are then released and analyzed via conventional means (e.g., microarray analysis, qRT-PCR, RNA sequencing, or Northern blot). While cross-linking-based ribonucleoprotein immunoprecipitation (RIP) has proven its utility within some contexts, it is technically challenging, inefficient, and suboptimal given the amount of time and resources (e.g., cells and antibodies) required. Additionally, these types of studies often require the use of over-expressed versions of proteins, which can introduce artifacts. Here, we describe a streamlined version of RIP that utilizes exclusion-based purification technologies. This approach requires significantly less starting material and resources compared to traditional RIP approaches, takes less time, which is tantamount given the labile nature of RNA, and can be used with endogenously expressed proteins. The method described here can be used to study RNA-protein interactions in a variety of cellular contexts.

Graphical abstract:

In the human cell cycle, complete replication of DNA is a fundamental process for the maintenance of genome integrity. Replication stress interfering with the progression of replication forks causes difficult-to-replicate regions to remain under-replicated until the onset of mitosis. In early mitosis, a homology-directed repair DNA synthesis, called mitotic DNA synthesis (MiDAS), is triggered to complete DNA replication. Here, we present a method to detect MiDAS in human U2OS 40-2-6 cells, in which repetitive lacO sequences integrated into the human chromosome evoke replication stress and concomitant incomplete replication of the lacO array. Immunostaining of BrdU and LacI proteins is applied for visualization of DNA synthesis in early mitosis and the lacO array, respectively. This protocol has been established to easily detect MiDAS at specific loci using only common immunostaining methods and may be optimized for the investigation of other difficult-to-replicate regions marked with site-specific binding proteins.

The in-cell western (ICW) is an immunocytochemical technique that has been used to screen for effects of siRNAs, drugs, and small molecule inhibitors. The reduced time and number of cells required to follow this protocol illustrates its semi-high-throughput nature. Performing a successful ICW protocol requires fixing and permeabilizing adherent cells directly in the plate that specifically exposes the epitope of interest. After blocking of non-specific proteins, the cells are incubated overnight with a primary antibody of interest, which is detected via a host-specific near-infrared fluorescently labeled LI-COR secondary antibody. In the final step, the plate is scanned using an Odyssey LI-COR Imaging System or similar, and each of the wells is quantified. For the first time, this technique has been demonstrated to be reproducibly utilized for semi-high-throughput selection of knockout or overexpression clones.

Graphical abstract:

Stable cell cloning is an essential aspect of biological research. All advanced genome editing tools rely heavily on stable, pure, single cell-derived clones of genetically engineered cells. For years, researchers have depended on single-cell dilutions seeded in 96- or 192-well plates, followed by microscopic exclusion of the wells seeded with more than or without a cell. This method is not just laborious, time-consuming, and uneconomical but also liable to unintentional error in identifying the wells seeded with a single cell. All these disadvantages may increase the time needed to generate a stable clone. Here, we report an easy-to-follow and straightforward method to conveniently create pure, stable clones in less than half the time traditionally required. Our approach utilizes cloning cylinders with non-toxic tissue-tek gel, commonly used for immobilizing tissues for sectioning, followed by trypsinization and screening of the genome-edited clones. Our approach uses minimal cell handling steps, thus decreasing the time invested in generating the pure clones effortlessly and economically.

Graphical abstract:

A schematic comparison showing the traditional dilution cloning and the method described here.

Here, a well-separated colony (in the green box) must be preferred over the colonies not well separated (in the red box).

Over the past years, research has made impressive breakthroughs towards the development and implementation of 3D cell models for a wide range of applications, such as drug development and testing, organogenesis, cancer biology, and personalized medicine. Opposed to 2D cell monolayer culture systems, advanced 3D cell models better represent the in vivo physiology. However, for these models to deliver scientific insights, appropriate investigation techniques are required. Despite the potential of fluorescence microscopy to visualize these models with high spatial resolution, sample preparation and imaging assays are not straightforward. Here, we provide different protocols of sample preparation for fluorescence imaging, for both matrix-embedded and matrix-free models (e.g., organoids and spheroids, respectively). Additionally, we provide detailed guidelines for imaging 3D cell models via confocal multi-photon fluorescence microscopy. We show that using these protocols, images of 3D cell culture systems can be obtained with sub-cellular resolution.

Graphical abstract:

Damage to the plasma membrane and loss of membrane integrity are detrimental to eukaryotic cells. It is, therefore, essential that cells possess an efficient membrane repair system to survive. However, the different cellular and molecular mechanisms behind plasma membrane repair have not been fully elucidated. Here, we present three complementary methods for plasma membrane wounding, and measurement of membrane repair and integrity. The first protocol is based on real time imaging of cell membrane repair kinetics in response to laser-induced injury. The second and third protocols are end point assays that provide a population-based measure of membrane integrity, after either mechanical injury by vortex mixing with glass beads, or by detergent-induced injury by digitonin in sublytic concentrations. The protocols can be applied to most adherent eukaryotic cells in culture, as well as cells in suspension.

The majority of biopsies in both basic research and translational cancer studies are preserved in the format of archived formalin-fixed paraffin-embedded (FFPE) samples. Profiling histone modifications in archived FFPE tissues is critically important to understand gene regulation in human disease. The required input for current genome-wide histone modification profiling studies from FFPE samples is either 10–20 tissue sections or whole tissue blocks, which prevents better resolved analyses. Nevertheless, it is desirable to consume a minimal amount of FFPE tissue sections in the analysis as clinical tissue of interest are limited. Here, we present FFPE tissue with antibody-guided chromatin tagmentation with sequencing (FACT-seq), highly sensitive method to efficiently profile histone modifications in FFPE tissue by combining a novel fusion protein of hyperactive Tn5 transposase and protein A (T7-pA-Tn5) transposition and T7 in vitro transcription. FACT-seq generates high-quality chromatin profiles from different histone modifications with low number of FFPE nuclei. We showed a very small piece of FFPE tissue section containing ~4000 nuclei is sufficient to decode H3K27ac modifications with FACT-seq. In archived FFPE human colorectal and human glioblastoma cancer tissue, H3K27ac FACT-seq revealed disease specific super enhancers. In summary, FACT-seq allows researchers to decode histone modifications like H3K27ac and H3K27me3 in archival FFPE tissues with high sensitivity, thus allowing us to understand epigenetic regulation.

Graphical abstract:

(i) FFPE tissue section; (ii) Isolated nuclei; (iii) Primary antibody, secondary antibody and T7-pA-Tn5 bind to targets; (iv) DNA purification; (v) In vitro transcription and sequencing library preparation; (vi) Sequencing