细胞生物学

The study of choroidal endothelial cells is essential for understanding the pathological mechanisms underlying choroidal neovascularization and other vision-threatening disorders. Traditional methods for isolating and culturing primary endothelial cells often yield mixed populations or require specialized equipment, limiting their widespread use. Here, we present a straightforward protocol for isolating and culturing primary mouse choroidal endothelial cells. This protocol involves enzymatic digestion of choroidal tissue, magnetic-activated cell sorting (MACS) to enrich CD31⁺ endothelial cells, and optimized culture conditions to promote cell proliferation and maintain endothelial phenotype. The protocol is strategic, reproducible, and requires minimal specialized equipment, making it accessible for researchers across various fields. By providing a robust method for obtaining pure choroidal endothelial cell cultures, this protocol facilitates the study of cell-specific behaviors and responses, advancing research into choroidal vascular diseases.

Since the establishment of the iSLK-BAC16 cell culture system, iSLK-BAC16 cells and their derivatives have been widely used for Kaposi’s sarcoma-associated herpesvirus (KSHV) studies. However, iSLK-BAC16 cells can be difficult to work with, in part due to the lack of standardized protocols and conflicting troubleshooting suggestions. Here, we describe the protocol for general iSLK-BAC16 cell culture and reactivation, which induces lytic KSHV replication and virion production. This protocol achieves robust levels of KSHV reactivation in our hands and can be readily used for studies of KSHV lytic infection mechanisms.

Corneal scarring, a significant cause of global blindness, results from various insults, including trauma, infections, and genetic disorders. The conventional treatment to replace scarred corneal tissues includes partial or full-thickness corneal transplantation using healthy donor corneas. However, only 1 in 70 individuals with treatable corneal scarring can undergo surgery, due to the limited supply of transplantable donor tissue. Our research focuses on cell-based strategies, specifically ex vivo–expanded corneal stromal stem cells (CSSCs), to address corneal scarring. Preclinical studies have demonstrated the efficacy of CSSC treatment in reducing corneal inflammation and fibrosis, inhibiting scar formation, and regenerating native stromal tissue. Mechanisms include CSSC differentiation into stromal keratocytes and the expression of regenerative cytokines. Here, we present a good manufacturing practice (GMP)-compliant protocol to isolate and expand human CSSCs. This method paves the way to produce clinical-grade CSSCs for transplantation and clinical trials.

Airway basal stem cells are the progenitor cells within the airway that exhibit the capacity to self-renew and give rise to multiple types of differentiated airway epithelial cells. This stem cell-derived epithelium displays organized architecture with functional attributes of the airway mucosa. A protocol has been developed to culture and expand human airway basal stem cells while preserving their stem cell properties and capacity for subsequent mucociliary differentiation. This achievement presents a previously unrealized opportunity to maintain a durable supply of progenitor cells derived from healthy donors to differentiate into human primary airway epithelium for cellular and molecular-based studies. Further, basal stem cells can be harvested from patients with a specific airway disease, such as cystic fibrosis, enabling investigation of potentially altered behavior of disease-specific cells in the appropriate context of the airway mucosa. Here we describe, in detail, a protocol for the serial expansion of airway basal stem cells to enable the generation of nearly unlimited airway basal cells that can be stored and readily available for subsequent culturing and differentiation. In addition, we describe culturing and differentiation of airway basal stem cells on permeable transwell filters at air-liquid interface to create functional mucociliary pseudostratified polarized airway epithelial mucosa.

The Microbial mutagenicity Ames test is a bacterial bioassay accomplished in vitro to evaluate the mutagenicity of various environmental carcinogens and toxins. While Ames test is used to identify the revert mutations which are present in strains, it can also be used to detect the mutagenicity of environmental samples such as drugs, dyes, reagents, cosmetics, waste water, pesticides and other substances which are easily solubilized in a liquid suspension. We present the protocol for conducting Ames test in the laboratory.

Networks of amyloid nanofibrils fabricated from common globular proteins such as lysozyme and β-lactoglobulin have material properties that mimic the extracellular microenvironment of many cell types. Cells cultured on such amyloid fibril networks show improved attachment, spreading and in the case of mesenchymal stem cells improved differentiation. Here we describe a detailed protocol for fabricating amyloid fibril networks suitable for eukaryotic cell culture applications.

Human cytomegalovirus (HCMV) infection is a major cause of morbidity and mortality in transplant patients and a leading cause of congenital birth defects (Saint Louis, 2016). Vaccination and therapeutic studies often require scalable cell culture production of wild type virus, represented by clinical isolates. Obtaining sufficient stocks of wild-type clinical HCMV is often labor intensive and inefficient due to low yield and genetic loss, presenting a barrier to studies of clinical isolates. Here we report a bioreactor method based on continuous infection, where retinal pigment epithelial (ARPE-19) cells adhered to microcarrier beads are infected in a bioreactor and used to produce high-titers of clinical isolate HCMV that maintain genetic integrity of key viral tropism factors and the viral genome. In this bioreactor, an end-stage infection can be maintained by regular addition of uninfected ARPE-19 cells, providing convenient preparation of 10⁷-10⁸ pfu/ml of concentrated TB40/E IE2-EYFP stocks without daily cell passaging or trypsinization. Overall, this represents a 100-fold increase in gain of virus production of 100-times compared to conventional static-culture plates, while requiring 90% less handling time. Moreover, this continuous infection environment has the potential to monitor infection dynamics with applications for real-time tracking of viral evolution.

This protocol aims at standardizing the procedure to obtain primary cultures of hippocampal and cerebrocortical neurons for in vitro experiments. Cultures should be prepared from cells isolated during embryonic development when neuronal precursor cells are not yet fully differentiated. This helps increasing the quality and quantity of cells, while offering minimal cell death that often occurs during dissociation of differentiated neurons. Cells plated under the appropriate conditions, either in Petri-dishes or in multi-well plates, will develop and establish synaptic contacts over time since the neuronal culture medium provides the nutrients and trophic factors required for differentiation. In this protocol we describe the methodology for the preparation of both cortical and hippocampal neuronal cultures.

A novel method to assess the dissolution of the core pluripotency transcription-factor circuit of mouse Embryonic Stem Cells (mESCs) has been developed (Ying et al., 2003; Betschinger et al., 2013). In order to efficiently identify genes essential for the break-down of the pluripotency network in mutant mESCs with proliferation defects, we adapted this ‘exit from pluripotency assay’ (Bodak et al., 2017; Cirera-Salinas et al., 2017). The protocol described here has been successfully applied to several mESC lines and is easily transposable from one laboratory to another.

A self-suppression mechanism of biofilm development in the cyanobacterium Synechococcus elongatus PCC 7942 was recently reported. These studies required quantification of biofilms formed by mutants impaired in the biofilm-inhibitory process. Here we describe in detail the use of chlorophyll measurements as a proxy for biomass accumulation in sessile and planktonic cells of biofilm-forming strains. These measurements allow quantification of the total biomass as estimated by chlorophyll level and representation of the extent of biofilm formation by depicting the relative fraction of chlorophyll in planktonic cells.