Methods in molecular biology最新文献

Enzyme assays are used to measure the activity or concentration of enzymes in biochemical or cell-based systems. Most enzyme assays are based on the detection of fluorescent, luminescent, or spectrophotometric endpoint signals. In recent years, they have been developed and widely used for diagnostics, mechanisms of action, and inflammatory activities. An enzyme assay essentially works by the conversion of a substrate into a product by the enzyme of interest. In this case, it is extremely important to know the optimal conditions for enzyme activity, as these affect the specificity and efficacy of the assay. For optimal reaction conditions, temperature, pH, and the presence of ions should be considered. In this chapter, the enzymatic assays for the detection of the enzymes N-acetylglucosaminidase (NAG), myeloperoxidase (MPO), and eosinophil peroxidase (EPO) are addressed. These assays are used to assess inflammatory parameters, for example, at the peripheral level in models of viral disease. They are based on an index of neutrophil, macrophage, or eosinophil accumulation in inflammatory tissues from animals by measuring the specific activity of the marker enzymes. The enzyme activity assays discussed here are based on colorimetric reactions compatible with any experimental model in which the respective cells has an active role. The advantage of using these enzymatic assays in inflammation response models is that they are simpler and less expensive compared to techniques such as Western blot or quantitative PCR.

Tight junctions, which are located on the apical side of epithelial cells, are key components of epithelial intercellular junctional complexes. Tight junctions seal the space between neighboring cells and act as a semipermeable barrier, preventing the paracellular transport of ions and molecules. The tight junctions are calcium-dependent as their disassembly can be triggered by the depletion of calcium ions, and the subsequent addition of calcium promotes the formation of tight junctions and the restoration of their barrier function. This reversible process, known as the calcium switch, is often used to study tight junction dynamics. This chapter describes the calcium switch protocol for disrupting and reestablishing tight junctions using MDCK cells as an in vitro model. It also provides protocols for evaluating tight junction formation and integrity using the noninvasive, quantitative transepithelial electrical resistance (TEER) assay.

Somatic hypermutation (SHM) is a critical process in adaptive immunity, enabling the generation of high-affinity antibodies through targeted mutations in immunoglobulin variable (IgV) regions. Here, we provide a comprehensive workflow combining immunization, molecular biology, and bioinformatics to investigate SHM mechanisms and outcomes. This is a general protocol for studying SHM at the VH186.2 region in C57BL/6 mice. While this method can be applied broadly, this chapter will detail the protocol used to test the effects of exonuclease 1(EXO1) knock-in mutation (Exo1^D173A, or Exo1^DA) or knock-out (KO) on hypermutation post-immunization by immunizing age-matched mice with NP(33)-CGG on alum. We start by immunizing and sacrificing the animals to obtain spleens for RNA extraction. We then create cDNA libraries and investigate VH186.2 region mutation to analyze SHM.

Clinical metagenomics (CMg) involves the untargeted sequencing of the genetic content of samples collected from patients and is a highly promising method for the diagnosis of infectious disease. Depending on the sample type, CMg can be reliant on the removal of the host genetic material from the sample to support detection of microbial pathogens, and this selective process (or an otherwise low abundance of microbial cells in the sample) may result in concentrations of DNA too low for productive sequencing. Whole genome amplification (WGA), the nonselective amplification of the total DNA of a sample, can be applied to significantly increase the concentration of DNA and enable CMg sequencing. This chapter describes the methods for the amplification of microbial DNA extracted from host-depleted wound swab samples using the GenomiPhi^™ V3 Ready-To-Go^™ (Cytiva) DNA WGA kit and host-depleted whole blood samples using the REPLI-g^® Single-Cell WGA kit (Qiagen). This is followed by the de-branching and bead-based clean-up of the amplified DNA, resulting in highly concentrated DNA ready for CMg DNA sequencing.

TNBC is an aggressive and metastatic subtype of breast cancer in which the TP53 mutation occurs frequently and is associated with particularly poor outcomes. Mutations in TP53 can disrupt the intrinsic function of the tumor suppressor as well as acquire oncogenic gain-of-function (GOF) activities. However, little is known about its oncogenic GOF mediators and functions. Targeted therapy for TNBC patients is thus one of the most urgent needs in breast cancer therapeutics, and identifying genes that have synthetic lethal interactions with mutant TP53 may be a promising approach. Sequential analysis of RNA-seq followed by high-throughput RNA interference screening (HTS-RNAi screening), as an intrinsic cellular mechanism for the identification of genes with synthetic lethality of mutant TP53, is a promising strategy for the treatment of mutant TP53 in TNBC and determining its impact on tumorigenesis.

CRISPR/Cas9-based genome editing is an inexpensive and efficient tool for genetic modification. Here, we present a methodological approach for establishing interleukin-17 receptor B (IL17RB) knockout cell lines using CRISPR/Cas9-mediated genomic deletion. The IL17RB gene encodes for a cytokine receptor that specifically binds to IL17B and IL17E and is overexpressed in various cancers. The method involves CRISPR design, CRISPR cloning, delivery of the CRISPR clone into cells, and verification of IL17RB gene deletion by deletion screening primer design, genomic DNA extraction, and polymerase chain reaction (PCR). A similar approach can be used for generating mammalian cell lines with gene knockout for other genes of interest.

Chimeric antigen receptor T cells (CAR T cells) therapy has revolutionarily changed the landscape of immunotherapy and been approved by the U. S Food and Drug Administration (FDA) since 2017 for several blood malignancies. To translate novel CAR T cells into clinical applications, it is essential to evaluate their antigen specificity, cytotoxic capacity, and off-target effects in vitro. A commonly used criteria to assess CAR T cell functionality involves detecting cytokine secretion following their engagement with target antigens. This chapter describes a method of combining intracellular cytokine staining and multi-color flow cytometry to measure CAR T cells activation following antigen stimulation.

Preimplantation genetic testing (PGT) is crucial for selecting embryos free of genetic abnormalities. However, existing PGT approaches often necessitate separate platforms for aneuploidy (PGT-A), monogenic disorders (PGT-M), and structural rearrangements (PGT-SR). This can drive up costs and operational complexity when multiple PGT tests are required for a single embryo. Here, we present a MALBAC-based method that integrates PGT-A, PGT-M, and PGT-SR into one unified platform.

TAPBPR has previously been identified as a homolog of tapasin, though the two proteins serve as mutually exclusive peptide editors. While tapasin functions solely as a constituent of the peptide loading complex, TAPBPR can function alone and independently of other chaperones and cofactors. An additional characteristic of TAPBPR is its lack of an endoplasmic-reticulum retention motif, which enables it to leak to the surface of particular cell types when overexpressed as well as an ability to promote peptide exchange at the cell surface as a recombinant soluble protein. The aforementioned features of TAPBPR provide the protein with unique capabilities for the characterization of its function, as well as the ability to dissect other properties of peptide loading such as peptide affinity for major histocompatibility complex class I and immune response to the presentation of immunoreactive peptide. Here, we describe the key methods used to decorate cells with peptides, permitting the assessment of the function of TAPBPR and its variants: peptide loading, peptide dissociation, and peptide exchange assays. The use of these assays confers the ability to dissect the catalytic function of TAPBPR and its variants, as well as conducting subsequent experiments utilizing the efficient decoration of cells with immunoreactive peptide.

Here, we describe a method to detect the oxidation of the pool of low molecular weight thiols (LMWT) in the infective stage of Trypanosoma brucei brucei in situ and non-invasively. The redox reporter cell line was generated by transfecting the parasites with a DNA construct coding for a redox-sensitive green fluorescent protein fused to human glutaredoxin 1 (hGrx1-roGFP2). The reporter gene is expressed in a tetracycline-inducible manner in the parasite's cytosol. roGFP2 displays a ratiometric and reversible change in fluorescence emission at 510 nm when excited at 405 and 488 nm, which is proportional to the changes in the ratio of oxidized vs. reduced LMWT trypanothione and glutathione. The role of hGrx1 is to catalyze a rapid equilibration of the LMWT pool with roGFP2 redox state, thereby allowing a biosensor response within seconds. The dynamic response of the biosensor enables the monitoring of cellular events in response to drugs or other stimuli in real time. The assay was adapted to a 96-well plate format for flow cytometry-based analysis. The fluorescent readout can be intensiometric or ratiometric, depending on the flow cytometer features, and the use of calibration controls is recommended for quantitative analysis. This bioassay can be applied to study fundamental questions of trypanosomatids' redox biology, as go/no-go criteria in drug discovery campaigns, and to investigate drug mode of action.