Cell Biology

Stem cell–based therapies have evolved to become a key component of regenerative medicine approaches to human pathologies. Exogenous stem cell transplantation takes advantage of the potential of stem cells to self-renew, differentiate, home to sites of injury, and sufficiently evade the immune system to remain viable for the release of anti-inflammatory cytokines, chemokines, and growth factors. Common to many pathologies is the exacerbation of inflammation at the injury site by proinflammatory macrophages. An increasing body of evidence has demonstrated that mesenchymal stromal cells (MSCs) can influence the immunophenotype and function of myeloid lineage cells to promote therapeutic effects. Understanding the degree to which MSCs can modulate the phenotype of macrophages within an inflammatory environment is of interest when considering strategies for targeted cell therapies. There is a critical need for potency assays to elucidate these intercellular interactions in vitro and provide insight into potential mechanisms of action attributable to the immunomodulatory and polarizing capacities of MSCs, as well as other cells with immunomodulatory potential. However, the complexity of the responses, in terms of cell phenotypes and characteristics, timing of these interactions, and the degree to which cell contact is involved, have made the study of these interactions challenging. To provide a research tool to study the direct interactions between MSCs and macrophages, we developed a potency assay that directly co-cultures MSCs with naïve macrophages under proinflammatory conditions. Using this assay, we demonstrated changes in the macrophage secretome and phenotype, which can be used to evaluate the abilities of the cell samples to influence the cell microenvironment. These results suggest the immunomodulatory effects of MSCs on macrophages while revealing key cytokines and phenotypic changes that may inform their efficacy as potential cellular therapies.

Key features

• The protocol uses monocytes differentiated into naïve macrophages, which are loosely adherent, have a relatively homogeneous genetic background, and resemble peripheral blood mononuclear cells–derived macrophages.

• The protocol requires a plate reader and a flow cytometer with the ability to detect six fluorophores.

• The protocol provides a quantitative measurement of co-culture conditions by the addition of a fixed number of freshly thawed or culture-rescued MSCs to macrophages.

• This protocol uses assessment of the secretome and cell harvest to independently verify the nature of the interactions between macrophages and MSCs.

Graphical overview

Proliferating cells need to cope with extensive cytoskeletal and nuclear remodeling as they prepare to divide. These events are tightly regulated by the nuclear translocation of the cyclin B1-CDK1 complex, that is partly dependent on nuclear tension. Standard experimental approaches do not allow the manipulation of forces acting on cells in a time-resolved manner. Here, we describe a protocol that enables dynamic mechanical manipulation of single cells with high spatial and temporal resolution and its application in the context of cell division. In addition, we also outline a method for the manipulation of substrate stiffness using polyacrylamide hydrogels. Finally, we describe a static cell confinement setup, which can be used to study the impact of prolonged mechanical stimulation in populations of cells.

Key features

• Protocol for microfabrication of confinement devices.

• Single-cell dynamic confinement coupled with high-resolution microscopy.

• Static cell confinement protocol that can be combined with super-resolution STED microscopy.

• Analysis of the mechanical control of mitotic entry in a time-resolved manner.

Graphical overview

The genome of the dengue virus codes for a single polypeptide that yields three structural and seven non-structural (NS) proteins upon post-translational modifications. Among them, NS protein-3 (NS3) possesses protease activity, involved in the processing of the self-polypeptide and in the cleavage of host proteins. Identification and analysis of such host proteins as substrates of this protease facilitate the development of specific drugs. In vitro cleavage analysis has been applied, which requires homogeneously purified components. However, the expression and purification of both S3 and erythroid differentiation regulatory factor 1 (EDRF1) are difficult and unsuccessful on many occasions. EDRF1 was identified as an interacting protein of dengue virus protease (NS3). The amino acid sequence analysis indicates the presence of NS3 cleavage sites in this protein. As EDRF1 is a high-molecular-weight (~138 kDa) protein, it is difficult to express and purify the complete protein. In this protocol, we clone the domain of the EDRF1 protein (C-terminal end) containing the cleavage site and the NS3 into two different eukaryotic expression vectors containing different tags. These recombinant vectors are co-transfected into mammalian cells. The cell lysate is subjected to SDS-PAGE followed by western blotting with anti-tag antibodies. Data suggest the disappearance of the EDRF1 band in the lane co-transfected along with NS3 protease but present in the lane transfected with only EDRF1, suggesting EDRF1 as a novel substrate of NS3 protease. This protocol is useful in identifying the substrates of viral-encoded proteases using ex vivo conditions. Further, this protocol can be used to screen anti-protease molecules.

Key features

• This protocol requires the cloning of protease and substrate into two different eukaryotic expression vectors with different tags.

• Involves the transfection and co-transfection of both the above recombinant vectors individually and together.

• Involves western blotting of the same PVDF membrane containing total proteins of the cell lysate with two different antibodies.

• Does not require purified proteins for the analysis of cleavage of any suspected substrate by the protease.

Graphical overview

The measurement of transepithelial electrical resistance across confluent cell monolayer systems is the most commonly used technique to study intestinal barrier development and integrity. Electric cell substrate impedance sensing (ECIS) is a real-time, label-free, impedance-based method used to study various cell behaviors such as cell growth, viability, migration, and barrier function in vitro. So far, the ECIS technology has exclusively been performed on cell lines. Organoids, however, are cultured from tissue-specific stem cells, which better recapitulate cell functions and the heterogeneity of the parent tissue than cell lines and are therefore more physiologically relevant for research and modeling of human diseases. In this protocol paper, we demonstrate that ECIS technology can be successfully applied on 2D monolayers generated from patient-derived intestinal organoids.

Key features

• We present a protocol that allows the assessment of various cell functions, such as proliferation and barrier formation, with ECIS on organoid-derived monolayers.

• The protocol facilitates intestinal barrier research on patient tissue-derived organoids, providing a valuable tool for disease modeling.

Autophagy is a conserved homeostatic mechanism involved in cellular homeostasis and many disease processes. Although it was first described in yeast cells undergoing starvation, we have learned over the years that autophagy gets activated in many stress conditions and during development and aging in mammalian cells. Understanding the fundamental mechanisms underlying autophagy effects can bring us closer to better insights into the pathogenesis of many disease conditions (e.g., cardiac muscle necrosis, Alzheimer’s disease, and chronic lung injury). Due to the complex and dynamic nature of the autophagic processes, many different techniques (e.g., western blotting, fluorescent labeling, and genetic modifications of key autophagy proteins) have been developed to delineate autophagy effects. Although these methods are valid, they are not well suited for the assessment of time-dependent autophagy kinetics. Here, we describe a novel approach: the use of DAPRed for autophagic flux measurement via live cell imaging, utilizing A549 cells, that can visualize and quantify autophagic flux in real time in single live cells. This approach is relatively straightforward in comparison to other experimental procedures and should be applicable to any in vitro cell/tissue models.

Key features

• Allows real-time qualitative imaging of autophagic flux at single-cell level.

• Primary cells and cell lines can also be utilized with this technique.

• Use of confocal microscopy allows visualization of autophagy without disturbing cellular functions.

The auxin-inducible degron (AID) system is a versatile tool in cell biology and genetics, enabling conditional protein regulation through auxin-induced degradation. Integrating CRISPR/Cas9 with AID expedites tagging and depletion of a required protein in human and mouse cells. The mechanism of AID involves interactions between receptors like TIR1 and the AID tag fused to the target protein. The presence of auxin triggers protein ubiquitination, leading to proteasome-mediated degradation. We have used AID to explore the mitotic functions of the replication licensing protein CDT1. Swift CDT1 degradation via AID upon auxin addition achieves precise mitotic inhibition, revealing defects in mitotic spindle structure and chromosome misalignment. Using live imaging, we found that mitosis-specific degradation of CDT1 delayed progression and chromosome mis-segregation. AID-mediated CDT1 inhibition surpasses siRNA-based methods, offering a robust approach to probe CDT1’s mitotic roles. The advantages of AID include targeted degradation and temporal control, facilitating rapid induction and reversal of degradation—contrasting siRNA’s delayed RNA degradation and protein turnover. In summary, the AID technique enhances precision, control, and efficiency in studying protein function and regulation across diverse cellular contexts. In this article, we provide a step-by-step methodology for generating an efficient AID-tagging system, keeping in mind the important considerations that need to be adopted to use it for investigating or characterizing protein function in a temporally controlled manner.

Key features

• The auxin-inducible degron (AID) system serves as a versatile tool, enabling conditional protein regulation through auxin-induced degradation in cell biology and genetics.

• Integration of CRISPR/Cas9 knock-in technology with AID expedites the tagging and depletion of essential proteins in mammalian cells.

• AID’s application extends to exploring the mitotic functions of the replication licensing protein CDT1, achieving precise mitotic inhibition and revealing spindle defects and chromosome misalignment.

• The AID system and its diverse applications advance the understanding of protein function and cellular processes, contributing to the study of protein regulation and function.

Graphical overview

Cdt1–auxin-inducible degron (AID) tagging workflow. (A) Schematic of the cloned Cdt1 gRNA vector and the repair template generated to endogenously tag the Cdt1 genomic locus with YFP and AID at the C-terminal using CRISPR/CAS9-based genome editing. The two plasmids are transfected into DLD1-TIR1 stable cells, followed by sorting and scaling up of YFP-positive single cells. (B) The molecular mechanism of auxin-induced proteasome-mediated degradation of the target protein (CDT1) shown at the bottom of the figure is well worked out.

Tears contain numerous secreted factors, enzymes, and proteins that help in maintaining the homeostatic condition of the eye and also protect it from the external environment. However, alterations to these enzymes and/or proteins during pathologies such as mechanical injury and viral or fungal infections can disrupt the normal ocular homeostasis, further contributing to disease development. Several tear film components have a significant role in curbing disease progression and promoting corneal regeneration. Additionally, several factors related to disease progression are secreted into the tear film, thereby serving as a valuable reservoir of biomarkers. Tears are readily available and can be collected via non-invasive techniques or simply from contact lenses. Tears can thus serve as a valuable and easy source for studying disease-specific biomarkers. Significant advancements have been made in recent years in the field of tear film proteomics, lipidomics, and transcriptomics to allow a better understanding of how tears can be utilized to gain insight into the etiology of diseases. These advancements have enabled us to study the pathophysiology of various disease states using tear samples. However, the mechanisms by which tears help to maintain corneal homeostasis and how they are able to form the first line of defense against pathogens remain poorly understood and warrant detailed in vitro studies. Herein, we have developed an in vitro assay to characterize the functional importance of patient isolated tears and their components on corneal epithelial cells. This novel approach closely mimics real physiological conditions and could help the researchers gain insight into the underlying mechanisms of ocular pathologies and develop new treatments.

Key features

• This method provides a new technique for analyzing the effect of tear components on human corneal epithelial cells.

• The components of the tears that are altered in response to diseases can be used as a biomarker for detecting ocular complications.

• This procedure can be further employed as an in vitro model for assessing the efficacy of drugs and discover potential therapeutic interventions.

Clearance of dying cells, named efferocytosis, is a pivotal function of professional phagocytes that impedes the accumulation of cell debris. Efferocytosis can be experimentally assessed by differentially tagging the target cells and professional phagocytes and analyzing by cell imaging or flow cytometry. Here, we describe an assay to evaluate the uptake of apoptotic cells (ACs) by human macrophages in vitro by labeling the different cells with commercially available dyes and analysis by flow cytometry. We detail the methods to prepare and label human macrophages and apoptotic lymphocytes and the in vitro approach to determine AC uptake. This protocol is based on previously published literature and allows for in vitro modeling of the efficiency of AC engulfment during continual efferocytosis process. Also, it can be modified to evaluate the clearance of different cell types by diverse professional phagocytes.

Graphical overview

Exosomes are a subpopulation of the heterogenous pool of extracellular vesicles that are secreted to the extracellular space. Exosomes have been purported to play a role in intercellular communication and have demonstrated utility as biomarkers for a variety of diseases. Despite broad interest in exosome biology, the conditions that regulate their secretion are incompletely understood. The goal of this procedure is to biochemically reconstitute exosome secretion in Streptolysin O (SLO)-permeabilized mammalian cells. This protocol describes the reconstitution of lyophilized SLO, preparation of cytosol and SLO-permeabilized cells, assembly of the biochemical reconstitution reaction, and quantification of exosome secretion using a sensitive luminescence-based assay. This biochemical reconstitution reaction can be utilized to characterize the molecular mechanisms by which different gene products regulate exosome secretion.

Key features

• This protocol establishes a functional in vitro system to reconstitute exosome secretion in permeabilized mammalian cells upon addition of cytosol, ATP, GTP, and calcium (Ca²⁺).

Graphical overview

This paper introduces simple analytical methods and bioassays to promptly assess the identity and function of in vitro cultured human Schwann cells (hSCs). A systematic approach is proposed to unequivocally discriminate hSCs from other glial cells, non-glial cells, and non-human SCs (authentication), identify hSCs at different stages of differentiation, and determine whether individual hSCs are proliferative or senescent. Examples of how to use distinct cell-based approaches for quality control and routine troubleshooting are provided to confirm the constitution (identity, purity, and heterogeneity) and potency (bioactivity) of hSC cultures from multiple sources. The bioassays are valuable for rapidly gauging the responses of hSCs to mitogenic and differentiating factors and ascertaining the cells’ basic properties before performing co-culture or cell grafting studies. The assays are image based and use adherent hSCs established in monoculture to simplify the experimental setup and interpretation of results. Finally, all sections contain thorough background information, notes, and references to facilitate decision making, data interpretation, and ad hoc method development for diverse applications.