Cell Biology

Patient-derived glioblastoma (GBM) cells are valuable models for GBM research due to their rarity and the highly lethal nature of this cancer. Preserving these cells through long-term cryopreservation is therefore essential for advancing future investigations. However, recent studies have reported that standard cell recovery protocols are inefficient, resulting in poor cell survival and limited regrowth. Here, we established an optimized culture protocol that enhances the recovery and expansion of patient-derived GBM cells by combining Matrigel with an increased concentration of fetal bovine serum (FBS). This approach significantly improves cell attachment and recovery after thawing cells that have been cryopreserved for more than a decade. Importantly, the recovered cells retain key phenotypic characteristics and remain suitable for downstream applications, including drug testing and spheroid formation. Together, this optimized protocol provides a novel strategy to increase the availability of patient-derived GBM cells by improving their efficient recovery from long-term cryopreservation, thereby maximizing their utility in GBM research.

Extracellular vesicles (EVs) are critical mediators of cell–cell communication and play a key role in male reproductive biology by modulating sperm function. This protocol describes a robust and reproducible workflow for isolating EVs from ram seminal plasma using size-exclusion chromatography (SEC) and assessing their uptake by ram spermatozoa. In contrast to ultracentrifugation-based methods, SEC provides a gentle and more efficient isolation approach that preserves EV integrity and functionality. A central innovation of this protocol is the use of carboxyfluorescein succinimidyl ester (CFSE)-labeled seminal plasma EVs (SP-EVs) to evaluate their incorporation into sperm cells through two complementary detection platforms: (i) flow cytometry with standard resolution and (ii) confocal microscopy, for spatial confirmation of EV–sperm interactions. By bridging the gap between EV isolation and functional analysis, this protocol provides a valuable tool for investigating the role of EV–cell interactions. Specifically, it offers potential applications in male fertility preservation, biomarker discovery, and the development of EV-based therapeutic strategies in reproductive medicine.

Organic solvent–based tissue clearing methods are widely used for whole-brain imaging but often compromise endogenous fluorescence. Existing protocols, such as iDISCO and fluorescence-preserving variants, have improved optical transparency but still present trade-offs between fluorescence retention, tissue stability, and workflow complexity. Here, we present MDISCO, a modified iDISCO-based clearing protocol designed to enhance preservation of endogenous fluorescence while maintaining high transparency and stable tissue morphology. MDISCO is directly compared with FDISCO+, an established fluorescence-preserving protocol, for the preservation of endogenous tdTomato and YFP. Performance across clearing steps is evaluated by measuring brain weight, anteroposterior and mediolateral dimensions, and optical transparency before and after solvent clearing and refractive index matching. Fluorescence preservation is assessed using whole-brain light-sheet microscopy with standardized imaging parameters to enable direct comparison. This protocol provides an accessible and high-throughput, reproducible workflow for solvent-based clearing with robust endogenous fluorescence preservation, offering clear advantages for whole-brain 3D imaging of genetically encoded fluorescent reporters.

Small ubiquitin-related modifiers (SUMOs) are covalently conjugated onto the proteome and serve as signaling molecules in many aspects of eukaryotic cell biology, from S. cerevisiae and C. elegans to H. sapiens. The conjugatable SUMO variants, SUMO1 and the almost identical SUMO2 and SUMO3 (designated SUMO2/3), are processed by an E1(SAE1:SAE2)-E2(UBC9)-E3 enzyme cascade to produce SUMO-modified proteins. The prerogative of the SUMO biology field is to identify and study the specific proteins undergoing SUMOylation, which grants us insights into the biological pathway of interest. This protocol was developed using the human osteosarcoma cell line U2OS to enable the investigation of SUMO conjugates in mitosis, the cell division phase of the cell cycle. We enrich the cell population for mitotic cells, which are isolated and subjected to stringent lysis conditions involving a high concentration of SDS and DTT in RIPA buffer, to promote complete protein denaturation. The lysates in high SDS RIPA buffer are diluted to reduce the overall SDS concentration and undergo conventional immunoprecipitation using SUMO1- or SUMO2/3-specific antibodies bound to protein A/G agarose beads. The samples are then compatible with downstream readouts such as western blots and mass spectrometry. This protocol detects endogenous SUMOylated proteins and avoids exogenous SUMO overexpression, which can alter SUMO conjugate formation. Furthermore, this denaturing protocol ensures only SUMOylated proteins are immunoprecipitated, and not their interactors.

Bovine muscle satellite cells (MuSC) and fibro-adipogenic progenitor cells (FAP) are muscle resident stem cells that are responsible for postnatal muscle growth, intramuscular fat deposition, and extracellular matrix generation. These cells are of increasing interest for the cultivated meat community due to their ability to generate all the major components of meat; additionally, these cells are of interest to conventional animal science research to elucidate mechanisms to improve meat quality. To use these cells for these goals, efficient and accurate cell isolation, culture, and differentiation are essential to evaluate their cell fate decisions and behaviors. In this protocol, we detail a simultaneous isolation of both MuSCs and FAPs with multiple intermediate stopping points, allowing for flexibility for day-of time constraints. We also detail improved growth conditions to maximize cell expansion and procedures to assess cell differentiation. This protocol provides a flexible isolation procedure that is compatible with sampling in modern slaughterhouses or from biopsies. Additionally, the differentiation procedures provide improved differentiation but still allow in vitro treatment and assessment.

Laccase2 (Lac2), a member of the phenoloxidase (PO) family, is an essential oxidase for melanin pigmentation in insects. The identification of the in vivo spatial distribution of Lac2 is crucial for understanding the molecular mechanisms underlying color pattern formation. However, it is technically difficult to determine the distribution because Lac2 expression peaks at late pupal stages, when adult cuticle sclerotization has already begun. Here, we report a simple and rapid protocol for estimating the distribution of endogenous PO proteins, prophenoloxidases (proPOs) and phenoloxidases (POs), in insect tissues. In this method, the spatial distribution of endogenous PO proteins is estimated based on staining patterns formed by dopamine melanin synthesis in tissues incubated in a solution containing isopropanol and dopamine. We validated that tissues collected at approximately 80% of the total pupal duration yielded staining patterns corresponding to adult melanin-forming regions in three insect species. By comparing staining patterns across developmental stages, this protocol enables estimation of the timing of color pattern formation. Furthermore, the contrast between stained and unstained regions within the same tissue allows region-specific sampling, thereby facilitating an investigation of the underlying molecular mechanisms regulating spatial PO distribution. Taken together, this method facilitates the study of melanin biosynthesis and enables the identification of the genes involved in regulating color pattern formation. This protocol does not require antibodies, transgenic lines, or specialized equipment and can be completed within a short time frame. Its effectiveness has been validated in multiple coleopteran and lepidopteran species, demonstrating its broad applicability as a versatile tool for studying insect pigmentation and color pattern formation.

Super-resolution imaging of synapses in intact brain tissue remains challenging because light scattering, photobleaching, and limited probe penetration, along with antigen accessibility within the densely packed postsynaptic densities (PSDs), constrain resolution and labeling efficiency. Here, we present a protocol utilizing thin brain cryosections and tau-stimulated emission depletion (STED) nanoscopy to visualize the intricate nano-architecture of excitatory synapses in situ. Slicing the brain into 6 μm sections allows for highly efficient and even penetration of probes throughout sections while ensuring that the resolution is not significantly impacted by the imaging depth of the tissue. We outline step-by-step instructions for labeling pre- and postsynaptic nano-architecture using antibodies and nanobodies, highlighting how fixative choice influences the labeling efficiency of synaptic proteins. While this protocol is compatible with both confocal and super-resolution imaging, when combined with rapid image acquisition times of tau-STED, it enables clear separation of key synaptic features in three dimensions with minimal photobleaching. Thus, this approach enables robust multiplex imaging of fluorescently labeled synaptic proteins in the brain, providing exceptional spatial resolution for visualization and quantification of synaptic nanoarchitecture in its native environment.

Single-cell RNA sequencing (scRNA-seq) is a powerful technique for exploring cellular heterogeneity and host–pathogen interactions. This protocol details the Zika virus (ZIKV)-targeted scRNA-seq workflow for preparing high-quality single-cell suspensions from the whole brain tissues of neonatal mice, high-quality single-cell sorting, cDNA reverse transcription, amplification, ZIKV enrichment and host transcriptome library preparation, and sequencing dataset integration in downstream analysis to complete the quantification of ZIKV RNA in individual cells.

Human tissue samples represent the gold standard for obtaining clinically relevant and translatable insight into disease processes that in vitro systems cannot fully reproduce. However, patient-derived samples are often limited in size and availability, limiting the number of downstream assays that can be performed. To maximize the use of invaluable human samples, we present a protocol for the tandem extraction of high-quality RNA and protein from the same tissue section. This method has been optimized for 15–30 mg tissue sections, enabling more experimental conditions and technical replicates, while minimizing intrasample variability associated with heterogeneous tissues. This protocol also avoids potentially hazardous solvents present in phenol-chloroform-based methods such as TRIzol, providing a safer and more accessible workflow without compromising biomolecule integrity. This protocol was developed and validated using atherosclerotic plaque tissue from carotid endarterectomy, a very challenging tissue type to work with due to extensive calcification, necrosis, and limited surgical availability. We have also validated this method using mouse aortic tissue and cultured THP-1 cells, demonstrating its versatility across sample input types. As this protocol relies on standard column-based RNA extraction kits and commonly available reagents for protein precipitation and extraction, this methodology is widely accessible and easy to implement as a standard, streamlined workflow.

Breast cancer (BC) is the most frequently diagnosed malignancy in women and a leading cause of cancer-related mortality worldwide. Current clinical management relies on molecular classification—based on estrogen receptor (ER), progesterone receptor (PR), HER2, and Ki67 expression—to guide prognosis and therapy. Triple-negative breast cancer (TNBC), which lacks ER, PR, and HER2 expression, represents 15%–20% of cases and is characterized by aggressive behavior, early recurrence, and a paucity of targeted treatment options. These challenges underscore the urgent need for improved preclinical models that better recapitulate tumor biology to accelerate therapeutic discovery. While conventional monolayer (2D) cultures have contributed significantly to cancer research, they fail to mimic critical features of the three-dimensional (3D) tumor microenvironment (TME), thereby limiting clinical translation. To address this gap, 3D spheroid models have emerged as a powerful intermediary, more accurately replicating in vivo conditions such as cell–cell and cell–matrix interactions, nutrient and oxygen gradients, and the development of hypoxic cores. These features make spheroids a physiologically relevant platform for studying complex processes like metastasis, drug resistance, and treatment response. Here, we present a robust, simple, and cost-effective protocol for generating uniform 3D spheroids. Our method enables consistent monitoring of spheroid formation and growth over time, with quantitative, image-based size analysis to ensure reproducibility and scalability. Designed for flexibility, the protocol is broadly applicable across diverse cell types, effectively bridging the gap between traditional 2D cultures and complex in vivo studies. By providing an accessible and reliable model of the 3D TME, this protocol opens new avenues for high-throughput drug screening, mechanistic studies of tumor progression, and the advancement of personalized medicine strategies in breast cancer and beyond.