Cell Biology

Following myocardial infarction (MI), myocardial cells undergo cell death, and the necrotic region is replaced by extracellular matrix (ECM) proteins such as collagens. Myofibroblasts are responsible for producing these ECM proteins. Cardiac myofibroblasts are differentiated from resident fibroblasts in response to inflammation. To date, genetically modified mice driven by the Periostin promoter and adeno-associated virus 9 (AAV9) carrying the Periostin promoter have been used for gene transfer into cardiac myofibroblasts. However, these methods require multiple steps and are time-consuming and expensive. Therefore, we developed a method for delivering genes into cardiac myofibroblasts using retroviruses. Specifically, the DNA of the target gene was transfected into Plat-E cells, which are packaging cells, to generate retroviruses. The virus-containing supernatant was then harvested, and the viruses were pelleted by centrifugation and suspended in PBS-containing polybrene. Subsequently, permanent occlusion of the left coronary artery was performed, and 20 μL of viral solution was immediately administered using a 29G needle at a position 1–2 mm below the ligation site in the heart of mice maintained in an open chest state. Using this method, we were able to introduce genes into the myofibroblasts of interest surrounding the MI site.

The antibody-uptake assay is a commonly used technique to monitor endocytosis of integral membrane proteins including transmembrane and glycosylphosphatidylinositol-anchored proteins (GPI-APs). The antibody-uptake assay typically involves incubating live cells with fluorophore-conjugated antibodies directed against the extracellular domain of the integral membrane protein of interest. Antibody uptake is then detected by flow cytometry or confocal microscopy. However, these detection modalities may be inaccessible to some labs or require extensive training to operate. Thus, we developed an easy and novel sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blot-based approach to the antibody-uptake assay that exploits the strong affinity between biotin and streptavidin. Instead of incubating cells with fluorophore-conjugated antibodies to monitor antibody uptake, our assay involves incubating cells with biotinylated antibodies, processing the cell lysates for western blot, and probing the membrane with detectably conjugated streptavidin. From preparation to quantification, this protocol requires less hands-on time than other approaches and is amenable to small-scale drug or siRNA screens. Here, we demonstrate the utility of our approach using the well-characterized misfolded GPI-AP, YFP-tagged C179A mutant of prion protein (YFP-PrP*), as our model substrate. YFP-PrP* constitutively traffics to the plasma membrane (PM), where it binds to anti-GFP antibody, and immediately undergoes endocytosis to lysosomes. To validate our protocol, we present measurements of antibody uptake under conditions known to enhance or inhibit YFP-PrP*’s traffic to the PM. Using this assay, we present new evidence that, under certain conditions, YFP-PrP* is able to undergo degradation via a pathway that does not involve exposure on the cell surface.

Nowadays, recombinant proteins are the focus of various research fields, and their use ranges from therapeutic investigations to cellular model systems for the development of therapeutic approaches. Cell systems used for the expression of recombinant proteins should be comparable in terms of yield and expression efficiency. In many research fields, it is desirable to obtain high protein concentrations. A method that combines an easy workflow with rapid results and affordable costs remains missing, and a standardized approach to determining protein concentration in transgenic cell lines is essential for more reliable data analysis. Our protocol demonstrates the cluster fluorescence-linked immunosorbent assay (FLISA), a technique that allows the exact quantification of comparable protein expression amounts. Moreover, it enables the detection of clustered or bound subunits of a protein without necessitating ultracentrifugation. In the present protocol, we demonstrate the utilization of two transgene cell lines, each expressing distinct recombinant proteins, to provide comparability of protein yields and detectable subunit clustering.

This protocol describes the isolation and flow cytometric analysis of extracellular vesicles (EVs) derived from red blood cells, endothelial cells, and platelets in human peripheral blood. The protocol includes steps for preparing platelet-free plasma, fluorescent antibody staining, gating strategies, and technical controls. This protocol was developed within a study on EV release in snakebite-associated thrombotic microangiopathy; the protocol addresses challenges such as variable autofluorescence and heterogeneity in EV origin. It is flexible and can be adapted for alternative antibody panels targeting different cell populations or EV subtypes, including leukocyte-derived EVs.

Most membrane and secreted proteins are transported from the endoplasmic reticulum (ER) to the Golgi apparatus and subsequently directed to their final destinations in the cell. However, the mechanisms underlying transport and cargo sorting remain unclear. Recent advancements in optical microscopy, combined with synchronized cargo protein release methods, have enabled the direct observation of cargo protein transport. We developed a new optically synchronized cargo release method called retention using the dark state of LOV2 (RudLOV). This innovative technique offers three exceptional control capabilities: spatial, temporal, and quantitative control of cargo release. RudLOV uses illumination to trigger transport and detect cargo. Consequently, the selection of an appropriate laser and filter set for controlling the illumination and/or detection is crucial. The protocol presented here provides step-by-step guidelines for obtaining high-resolution live imaging data using RudLOV, thereby enabling researchers to investigate intracellular cargo transport with unprecedented precision and control.

Autophagy plays a crucial role in cellular homeostasis and is responsible for removing and degrading damaged cytoplasmic cargo. This lysosome-mediated catabolic process removes defective organelles and misfolded proteins, and impaired autophagy has been directly linked to ageing and numerous diseases. This emphasises the importance of developing intervention methods to counteract this dysregulation. One promising intervention is thermal therapy, specifically hyperthermia, which is described in this protocol. In order to investigate this form of treatment, a rapid and reliable detection method is required to allow comparison of autophagy status under different conditions. While methods such as transmission electron microscopy (TEM) or western blotting can provide valuable structural analysis, they are often time-consuming and expensive, and are not suitable for small, round cells such as peripheral blood mononuclear cells (PBMCs). The method described in this protocol enables absolute quantification of PBMCs using the Guava^® Autophagy Detection kit after heat treatment with water-filtered infrared-A radiation (wIRA), compared with an untreated control. This method is based on antibody labelling, and subsequent flow cytometric analysis enables the number of autophagosomes to be determined by measuring the FITC intensity. This protocol provides rapid, reliable results and can be adapted to investigate not only heat therapy, but also other interventions, such as caloric restriction.

Adult muscle stem cells (MuSCs) are the key cellular source for regenerating skeletal muscle in vertebrates. MuSCs are typically identified in skeletal muscle by the expression of the paired box protein 7 (PAX7) protein. Here, we developed a combined RNA fluorescent in situ hybridization (FISH) using RNAscope technology and an immunofluorescence (IF) protocol for the simultaneous detection of Pax7 mRNA and PAX7 protein in individual MuSCs in vivo. Interestingly, we show that while most PAX7⁺ (protein) MuSCs express Pax7 mRNA, there is a subset of Pax7⁺ (mRNA) cells that do not express PAX7 protein. Altogether, we developed a combined FISH/IF protocol that allows for the co-detection of mRNA and protein in MuSCs in vivo, a strategy that can be applied to any target gene. The functional significance of the Pax7-expressing subset of cells lacking PAX7 protein prior to injury remains unknown.

Chemically induced murine colitis models are widely used to understand intestinal homeostasis and inflammatory responses during acute and chronic gut inflammation, such as inflammatory bowel disease (IBD). Resident populations of immune cells, together with those recruited during an inflammatory response, maintain intestinal immunity by mounting an effective immune response to enteropathogenic microbes while at the same time maintaining tolerance against commensals. To better understand the disease mechanism, studying different immune cell populations and their dynamic changes during infection and inflammation is essential. However, isolating healthy and viable immune populations, particularly hyperactivated neutrophils and macrophages from the inflamed gut (i.e., active disease site), is challenging as tissues are usually subjected to rigorous enzymatic digestion for an extended period. Here, we describe a method that uses a cell dissociator (Medimachine II from Syntec International) to separate intestinal tissue after short enzymatic digestion to obtain a single-cell suspension. This technique facilitates the isolation of immune cells from mouse intestinal tissues in high quantity and with superior viability in a very short time frame. This protocol delivers 80%–90% cell viability, which is 1.5 to 2-fold higher than conventional methods of isolating cells from inflamed mouse colons. The composition, phenotype, activation state, and gene expression profile of cells isolated using this protocol can be assessed by using multiple methods, including, but not limited to, flow cytometry, quantitative PCR, immunoblotting, mass spectrometry, single-cell RNA sequencing, and functional readouts such as reactive oxygen species (ROS) production.

This protocol describes an ex vivo co-culture method to assess CD8⁺ T-cell activation, proliferation, and cytotoxic potential using bulk splenocytes isolated from immunocompetent mice. Mouse splenocytes are stimulated with anti-CD3 and anti-CD28 antibodies to activate CD8⁺ T cells, which are then co-incubated with either cancer cells or cancer cell–derived conditioned media (CM) to evaluate tumor-driven modulation of immune cell functions. The use of unfractionated splenocytes preserves physiological cell–cell interactions, eliminating the need for exogenous interleukin (IL-2) and bypassing flow sorting, which simplifies the workflow and reduces experimental variability. CD8⁺ T-cell responses are measured via flow cytometry, using markers of proliferation (CFSE dilution), activation (CD69), and effector function (Granzyme B and IFNγ). Additionally, immune-mediated tumor cell death is evaluated by Annexin-V/7-AAD staining. Together, this experimental platform supports the investigation of both cell contact-dependent and contact-independent mechanisms of immune cell modulation in a cost-effective and reproducible setting.

Studying G protein-coupled receptor (GPCR) activation of heterotrimeric G proteins is crucial for understanding diverse physiological processes and developing novel therapeutics. Traditional methods to assay GPCR activation of G proteins, including assays of second messengers and biosensors, involve complex or indirect procedures. However, second messengers like cAMP and calcium are not direct readouts of GPCR activity due to signaling crosstalk, while biosensors can have undesired consequences due to structural alteration caused by fluorescent protein insertion. Here, we present a streamlined protocol employing GST-tagged bait proteins and epitope-embedded Gα subunits to achieve direct monitoring of Gα activity within cells. This method involves purification of GST-tagged bait constructs from bacteria and subsequent direct interaction studies with GluGlu-tagged Gα proteins expressed in any human cells of interest by including GST-tagged bait proteins in the cell lysis buffer. The approach enables sensitive detection of activated Gα within cells following extracellular stimulation. Advantages of this protocol include high sensitivity, enhanced monitoring of GPCR signaling dynamics under physiologically relevant conditions with minimum alteration in Gα, and the ability to distinguish between highly homologous isoforms within the same Gα family.