Cell Biology

When plants undergo senescence or experience carbon starvation, leaf cells degrade proteins in the chloroplasts on a massive scale via autophagy, an evolutionarily conserved process in which intracellular components are transported to the vacuole for degradation to facilitate nutrient recycling. Nonetheless, how portions of chloroplasts are released from the main chloroplast body and mobilized to the vacuole remains unclear. Here, we developed a method to observe the autophagic transport of chloroplast proteins in real time using confocal laser-scanning microscopy on transgenic plants expressing fluorescently labeled chloroplast components and autophagy-associated membranes. This protocol enabled us to track changes in chloroplast morphology during chloroplast-targeted autophagy on a timescale of seconds, and it could be adapted to monitor the dynamics of other intracellular processes in plant leaves.

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.

Cellular phenomena such as signal integration and transmission are based on the correct spatiotemporal organization of biomolecules within the cell. Therefore, the targeted manipulation of such processes requires tools that can precisely induce the localizations and interactions of the key players relevant to these processes with high temporal resolution. Chemically induced dimerization (CID) techniques offer a powerful means to manipulate protein function with high temporal resolution and subcellular specificity, enabling direct control over cellular behavior. Here, we present the detailed synthesis and application of dual SLIPT and dual SLIPT^NVOC, which expand the SLIPT (self-localizing ligand-induced protein translocation) platform by incorporating a dual-ligand CID system. Dual SLIPT and dual SLIPT^NVOC independently sort into the inner leaflet of the plasma membrane via a lipid-like anchoring motif, where they present the two headgroup moieties trimethoprim (TMP) and HaloTag ligand (HTL), which recruit and dimerize any two ^iK6eDHFR- and HOB-tagged proteins of interest (POIs). Dual-SLIPT^NVOC furthermore enables this protein dimerization of POIs at the inner leaflet of the plasma membrane in a pre-determined order and light-controlled manner. In this protocol, we detail the synthetic strategy to access dual SLIPT and dual SLIPT^NVOC, while also providing the underlying rationale for key design and synthetic decisions, with the aim of offering a streamlined, accessible, and broadly implementable methodology. In addition to the detailed synthesis, we present representative applications and typical experimental outcomes and recommend strategies for data analysis to support effective use of the system. Notably, dual SLIPT and dual SLIPT^NVOC represent the first CID systems to emulate endogenous lipidation-driven membrane targeting, while retaining hallmark advantages of CID technology—the precision over POI identity, recruitment sequence, high spatiotemporal control, and “plug-and-play” flexibility.

Three-dimensional (3D) human brain tissue models derived from induced pluripotent stem cells (iPSCs) have transformed the study of neural development and disease in vitro. While cerebral organoids offer high structural complexity, their large size often leads to necrotic core formation, limiting reproducibility and challenging the integration of microglia. Here, we present a detailed, reproducible protocol for generating multi-cell type 3D neurospheres that incorporate neurons, astrocytes, and optionally microglia, all derived from the same iPSCs. While neurons and astrocytes differentiate spontaneously from neural precursor cells, generated by dual SMAD-inhibition (blocking BMP and TGF-b signaling), microglia are generated in parallel and can infiltrate the mature neurosphere tissue after plating neurospheres into 48-well plates. The system supports a range of downstream applications, including functional confocal live imaging of GCaMP6f after adeno-associated virus (AAV) transduction of neurospheres or immunofluorescence staining after fixation. Our approach has been successfully implemented across multiple laboratories, demonstrating its robustness and translational potential for studying neuron–glia interactions and modeling neurodegenerative processes.

Extracellular vesicles (EVs) have emerged as promising carriers for the targeted delivery of therapeutic proteins to specific cells. Previously, we demonstrated that genetically engineered EVs can be used for targeted protein delivery. This protocol details the generation of mannose receptor (CD206)-targeted EVs using a modular plasmid system optimized for production in HEK293T cells. Three plasmids enable customizable EV budding, cargo loading, and surface modification for targeting to antigen-presenting cells (APCs). EVs are isolated via differential centrifugation and chromatography, characterized using transmission electron microscopy (TEM) and nanoparticle tracking analysis (NTA), and validated through functional uptake assays in primary human activated dendritic cells. Our approach combines flexibility in engineering required EVs with robust, reproducible isolation and characterization workflows. Its modularity allows easy adaptation to alternative targets or cargoes, which can be validated immediately through in vitro testing.

Accurate labeling of excitatory postsynaptic sites remains a major challenge for high-resolution imaging due to the dense and sterically restricted environment of the postsynaptic density (PSD). Here, we present a protocol utilizing Sylites, 3 kDa synthetic peptide probes that bind with nanomolar affinity to key postsynaptic markers, PSD-95 and Gephyrin. eSylites (excitatory Sylites) specifically target the PDZ1 and PDZ2 domains of PSD-95, enabling precise and efficient labeling of excitatory postsynaptic density (ePSD). In contrast, iSylites (inhibitory Sylites) bind to the dimerizing E-domain of the Gephyrin C-terminus, allowing selective visualization of inhibitory postsynaptic density (iPSD). Their small size reduces linkage error and enhances accessibility compared to conventional antibodies, enabling clear separation of PSD-95 nanodomains in super-resolution microscopy. The protocol is compatible with co-labeling using standard antibodies and integrates seamlessly into multichannel immunocytochemistry workflows for primary neurons and brain tissue. This method enables robust, reproducible labeling of excitatory synapses with enhanced spatial resolution and can be readily adapted for expansion microscopy or live-cell applications.

Translation is a key step in decoding the genetic information stored in DNA. Regulation of translation is an important step in gene expression control and is essential for healthy organismal development and behavior. Despite the importance of translation regulation, its impact and dynamics remain only partially understood. One reason is the lack of methods that enable the real-time visualization of translation in the context of multicellular organisms. To overcome this critical gap, microscopy-based methods that allow visualization of translation on single mRNAs in living cells and animals have been developed. A powerful approach is the SunTag system, which enables real-time imaging of nascent peptide synthesis with high spatial and temporal resolution. This protocol describes the implementation and use of the SunTag translation imaging system in the small round worm Caenorhabditis elegans. The protocol provides details on how to design, carry out, and interpret experiments to image translation dynamics of an mRNA of interest in a cell type of choice of living C. elegans. The ability to image translation live enables better understanding of translation and reveals the mechanisms underlying the dynamics of cell type–specific and subcellular localization of translation in development.

Adipose tissue macrophages (ATMs) critically influence obesity-induced inflammation and metabolic dysfunction. Recent studies identified distinct ATM subsets characterized by markers such as CD11c, CD9, and Trem2, associated with pro-inflammatory and metabolically activated states. This protocol outlines a detailed, reproducible methodology for isolating, characterizing, and sorting these ATM subsets from murine epididymal white adipose tissue (eWAT) using multicolor flow cytometry. Key steps include stromal vascular fraction (SVF) isolation, immunophenotyping, sequential gating strategies, and fluorescence-activated cell sorting (FACS) for downstream gene expression analysis. The protocol was validated in diet-induced obese (DIO) mice treated with the IRE1 RNase inhibitor STF-083010, demonstrating its utility for studying ATMs in the context of obesity and metabolic disease.

Diabetes lacks concrete curative strategies due to diverse aetiologies and, therefore, represents the perfect candidate for cell replacement therapy, since it is caused by either an absolute (type 1 diabetes) or relative (type 2 diabetes) defect in the insulin-producing beta cells of the pancreas. Pancreatic alpha cells are a promising source for transdifferentiation into insulin-producing cells as they share a common developmental origin with beta cells and exhibit a certain degree of cellular plasticity. Furthermore, impairment of glucagon signaling in diabetes leads to a marked increase in alpha cell mass, raising the possibility that such alpha cell hyperplasia provides an increased supply of alpha cells for their transdifferentiation into new beta cells.

In this protocol, we used the modular epigenetic CRISPR/dCas9 toolbox for targeted DNA methylation (EpiCRISPR) and silencing of the Arx gene (Aristaless Related Homeobox, Arx), which is essential for the maintenance of alpha cell identity. Methylation-based silencing of Arx initiates the reprogramming of pancreatic alpha cells into insulin-producing cells. As a key novelty, this protocol provides a direct route for epigenetically induced transdifferentiation of mouse pancreatic alpha TC1-6 cells into insulin-producing cells and thereby confirms a proof of concept of reversible cellular epigenetic reprogramming in vitro. In addition, this streamlined workflow addresses the inherent challenges of transfecting clustered alpha TC1-6 cells by optimizing their dissociation into single-cell suspensions, thereby improving uptake and reproducibility.

In summary, this approach for cell transdifferentiation involves precise epigenetic editing of a lineage-specific marker gene, thereby enabling direct lineage conversion in a safe and versatile strategy to generate insulin-producing cells by epigenetic reprogramming. In contrast to approaches that rely on viral vectors or permanent genome editing, this method reduces the risk of off-target effects and immunogenic responses while ensuring reproducibility. The combination of efficiency and precision makes it a valuable tool to advance regenerative approaches for diabetes therapy and to explore the epigenetic regulation of cell identity.