Cancer 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.

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.

Immunohistochemistry (IHC) and immunofluorescence (IF) are fundamental molecular biology techniques to assess protein expression. However, the melanin present normally in the eye in the uveal tract (choroid, iris, and ciliary body) and the retinal pigment epithelium (RPE) poses a significant challenge for IHC and IF. This is because melanin interferes with both chromogenic and fluorescent detection methods. Additionally, formalin fixation, which is commonly used for IHC, can result in shrinkage and loss of cellular detail in the eye. This protocol provides an optimized approach using Davidson’s fixative with a hydrogen peroxide bleaching step to eliminate melanin interference in the mouse eye, improving the quality and interpretability of IHC analyses of the uveal tract and RPE. It is particularly useful for the analysis of uveal melanoma.

Formalin-fixed paraffin-embedded (FFPE) slides are essential for histological and immunohistochemical analyses of organoids. Conventional preparation of FFPE slides from organoids embedded in basement membrane extract (BME) presents several challenges. During the fixation step, dehydration often causes collapse of the BME, which normally supports the three-dimensional architecture of organoids. As a result, organoids may lose their original morphology, particularly in the case of cystic or structurally delicate types, leading to distortion and reduced reliability in downstream histological evaluation. Here, we introduce a straightforward protocol that improves the reliability of FFPE slide preparation for BME-based organoids by enhancing sample integrity and sectioning quality. By using 2% agarose as a mold during the embedding process, organoids grown in BME were effectively stabilized, enabling reliable preservation of their morphology throughout FFPE slide preparation. This method effectively addresses the difficulties in processing structurally delicate organoids and allows robust preparation of diverse cancer organoid morphologies—such as cystic, dense, and grape-like structures—while maintaining their native three-dimensional architecture. Our approach simplified the technical process while ensuring reliable histopathological analysis, making it a valuable tool for cancer research and personalized medicine.

In the human cell cycle, complete replication of DNA is a fundamental process for the maintenance of genome integrity. Replication stress interfering with the progression of replication forks causes difficult-to-replicate regions to remain under-replicated until the onset of mitosis. In early mitosis, a homology-directed repair DNA synthesis, called mitotic DNA synthesis (MiDAS), is triggered to complete DNA replication. Here, we present a method to detect MiDAS in human U2OS 40-2-6 cells, in which repetitive lacO sequences integrated into the human chromosome evoke replication stress and concomitant incomplete replication of the lacO array. Immunostaining of BrdU and LacI proteins is applied for visualization of DNA synthesis in early mitosis and the lacO array, respectively. This protocol has been established to easily detect MiDAS at specific loci using only common immunostaining methods and may be optimized for the investigation of other difficult-to-replicate regions marked with site-specific binding proteins.

Stable cell cloning is an essential aspect of biological research. All advanced genome editing tools rely heavily on stable, pure, single cell-derived clones of genetically engineered cells. For years, researchers have depended on single-cell dilutions seeded in 96- or 192-well plates, followed by microscopic exclusion of the wells seeded with more than or without a cell. This method is not just laborious, time-consuming, and uneconomical but also liable to unintentional error in identifying the wells seeded with a single cell. All these disadvantages may increase the time needed to generate a stable clone. Here, we report an easy-to-follow and straightforward method to conveniently create pure, stable clones in less than half the time traditionally required. Our approach utilizes cloning cylinders with non-toxic tissue-tek gel, commonly used for immobilizing tissues for sectioning, followed by trypsinization and screening of the genome-edited clones. Our approach uses minimal cell handling steps, thus decreasing the time invested in generating the pure clones effortlessly and economically.

Graphical abstract:

A schematic comparison showing the traditional dilution cloning and the method described here. Here, a well-separated colony (in the green box) must be preferred over the colonies not well separated (in the red box).

Over the past years, research has made impressive breakthroughs towards the development and implementation of 3D cell models for a wide range of applications, such as drug development and testing, organogenesis, cancer biology, and personalized medicine. Opposed to 2D cell monolayer culture systems, advanced 3D cell models better represent the in vivo physiology. However, for these models to deliver scientific insights, appropriate investigation techniques are required. Despite the potential of fluorescence microscopy to visualize these models with high spatial resolution, sample preparation and imaging assays are not straightforward. Here, we provide different protocols of sample preparation for fluorescence imaging, for both matrix-embedded and matrix-free models (e.g., organoids and spheroids, respectively). Additionally, we provide detailed guidelines for imaging 3D cell models via confocal multi-photon fluorescence microscopy. We show that using these protocols, images of 3D cell culture systems can be obtained with sub-cellular resolution.

Graphical abstract:

Current methods to obtain mesenchymal stem cells (MSCs) involve sampling, culturing, and expanding of primary MSCs from adipose, bone marrow, and umbilical cord tissues. However, the drawbacks are the limited numbers of total cells in MSC pools, and their decaying stemness during in vitro expansion. As an alternative resource, recent ceiling culture methods allow the generation of dedifferentiated fat cells (DFATs) from mature adipocytes. Nevertheless, this process of spontaneous dedifferentiation of mature adipocytes is laborious and time-consuming. This paper describes a modified protocol for in vitro dedifferentiation of adipocytes by employing an additional physical stimulation, which takes advantage of augmenting the stemness-related Wnt/β-catenin signaling. Specifically, this protocol utilizes a polyethylene glycol (PEG)-containing hypertonic medium to introduce extracellular physical stimulation to obtain higher efficiency and introduce a simpler procedure for adipocyte dedifferentiation.

Connexins are membrane bound proteins that facilitate direct and local paracrine mediated cell-to-cell communication through their ability to oligomerise into hexameric hemichannels. When neighbouring channels align, they form gap-junctions that provide a direct route for information transfer between cells. In contrast to intact gap junctions, which typically open under physiological conditions, undocked hemichannels have a low open probability and mainly open in response to injury. Hemichannels permit the release of small molecules and ions (approximately 1kDa) into the local intercellular environment, and excessive expression/activity has been linked to a number of disease conditions. Carboxyfluorescein dye uptake measures functional expression of hemichannels, where increased hemichannel activity/function reflects increased loading. The technique relies on the uptake of a membrane-impermeable fluorescent tracer through open hemichannels, and can be used to compare channel activity between cell monolayers cultured under different conditions, e.g. control versus disease. Other techniques, such as biotinylation and electrophysiology can measure cell surface expression and hemichannel open probability respectively, however, carboxyfluorescein uptake provides a simple, rapid and cost-effective method to determine hemichannel activity in vitro in multiple cell types.

Graphic abstract


Using dye uptake to measure hemichannel activity

The in vitro cell adhesion assay is a quantitative method for measuring selective cell adhesion to specific proteins. Traditionally, cell adhesion assays employ purified protein immobilized on a solid glass or plastic surface. Here, we describe a transient 293T cell transfection-based cell adhesion assay to study selective cell adhesion of a specific cell type to a protein of interest. In this protocol, 293T cells are transfected with a mammalian expression plasmid containing mSiglec1 cDNA or an empty plasmid as a mock control and are then cultured to form a monolayer. Subsequently, these Siglec1-expressing and mock-transfected 293T cell monolayers are used for cell adhesion assays with GFP-expressing B16F10 cells. The number of GFP⁺ cancer cells adhering to each 293T monolayer is a quantitative mean to compare the selective adhesiveness of cancer cells to Siglec1. This method eliminates the need to express and purify the protein of interest to perform in vitro cell adhesion assays and can easily be performed with difficult-to-purify proteins while maintaining their native in situ structure.