Cell Biology

The cation-independent mannose 6-phosphate receptors (CI-M6PR) bind newly synthesized mannose 6-phosphate (Man-6-P)-tagged enzymes in the Golgi and transport them to late endosomes/lysosomes, providing them with degradative functions. Following the cargo delivery, empty receptors are recycled via early/recycling endosomes back to the trans-Golgi network (TGN) retrogradely in a dynein-dependent motion. One of the most widely used methods for studying the retrograde trafficking of CI-M6PR involves employing the CD8α-CI-M6PR chimera. This chimera, comprising a CD8 ectodomain fused with the cytoplasmic tail of the CI-M6PR receptor, allows for labeling at the plasma membrane, followed by trafficking only in a retrograde direction. Previous studies utilizing the CD8α-CI-M6PR chimera have focused mainly on colocalization studies with various endocytic markers under steady-state conditions. This protocol extends the application of the CD8α-CI-M6PR chimera to live cell imaging, followed by a quantitative analysis of its motion towards the Golgi. Additionally, we present an approach to quantify parameters such as speed and track lengths associated with the motility of CD8α-CI-M6PR endosomes using the Fiji plugin TrackMate.

Apolipoprotein B (APOB) is the primary structural protein of atherogenic lipoproteins, which drive atherogenesis and thereby lead to deadly cardiovascular diseases (CVDs). Plasma levels of APOB-containing lipoproteins are tightly modulated by LDL receptor–mediated endocytic trafficking and cargo receptor–initiated exocytic route; the latter is much less well understood. This protocol aims to present an uncomplicated yet effective method for detecting APOB/lipoprotein secretion. We perform primary mouse hepatocyte isolation and culture coupled with well-established techniques such as immunoblotting for highly sensitive, specific, and semi-quantitative analysis of the lipoprotein secretion process. Its inherent simplicity facilitates ease of operation, rendering it a valuable tool widely utilized to explore the intricate landscape of cellular lipid metabolism and unravel the mechanistic complexities underlying lipoprotein-related diseases.

Various protocols have been proven effective in the directed differentiation of mouse and human pluripotent stem cells into skeletal muscles and used to study myogenesis. Current 2D myogenic differentiation protocols can mimic muscle development and its alteration under pathological conditions such as muscular dystrophies. 3D skeletal muscle differentiation approaches can, in addition, model the interaction between the various cell types within the developing organoid. Our protocol ensures the differentiation of human embryonic/induced pluripotent stem cells (hESC/hiPSC) into skeletal muscle organoids (SMO) via cells with paraxial mesoderm and neuromesodermal progenitors’ identity and further production of organized structures of the neural plate margin and the dermomyotome. Continuous culturing omits neural lineage differentiation and promotes fetal myogenesis, including the maturation of fibroadipogenic progenitors and PAX7-positive myogenic progenitors. The PAX7 progenitors resemble the late fetal stages of human development and, based on single-cell transcriptomic profiling, cluster close to adult satellite cells of primary muscles. To overcome the limited availability of muscle biopsies from patients with muscular dystrophy during disease progression, we propose to use the SMO system, which delivers a stable population of skeletal muscle progenitors from patient-specific iPSCs to investigate human myogenesis in healthy and diseased conditions.

Plasma membrane proteins mediate important aspects of physiology, including nutrient acquisition, cell–cell interactions, and monitoring homeostasis. The trafficking of these proteins, involving internalisation from and/or recycling back to the cell surface, is often critical to their functions. These processes can vary among different proteins and cell types and states and are still being elucidated. Current strategies to measure surface protein internalisation and recycling are typically microscopy or biochemical assays; these are accurate but generally limited to analysing a homogenous cell population and are often low throughput. Here, we present flow cytometry–based methods involving probe-conjugated antibodies that enable quantification of internalisation or recycling rates at the single-cell level in complex samples. To measure internalisation, we detail an assay where the protein of interest is labelled with a specific antibody conjugated to a fluorescent oligonucleotide-labelled probe. To measure recycling, a specific antibody conjugated to a cleavable biotin group is employed. These probes permit the differentiation of molecules that have been internalised or recycled from those that have not. When combined with cell-specific marker panels, these methods allow the quantitative study of plasma membrane protein trafficking dynamics in a heterogenous cell mixture at the single-cell level.

Precision-cut lung slices (PCLS), ex vivo 3D lung tissue models, have been widely used for various applications in lung research. PCLS serve as an excellent intermediary between in vitro and in vivo models because they retain all resident cell types within their natural niche while preserving the extracellular matrix environment. This protocol describes the TReATS (TAT-Cre recombinase-mediated floxed allele modification in tissue slices) method that enables rapid and efficient gene modification in PCLS derived from adult floxed animals. Here, we present detailed protocols for the TReATS method, consisting of two simple steps: PCLS generation and incubation in a TAT-Cre recombinase solution. Subsequent validation of gene modification involves live staining and imaging of PCLS, quantitative real-time PCR, and cell viability assessment. This four-day protocol eliminates the need for complex Cre-breeding, circumvents issues with premature lethality related to gene mutation, and significantly reduces the use of animals. The TReATS method offers a simple and reproducible solution for gene modification in complex ex vivo tissue-based models, accelerating the study of gene function, disease mechanisms, and the discovery of drug targets.

Cultured mammalian cells are a common model system for the study of epithelial biology and mechanics. Epithelia are often considered as pseudo–two dimensional and thus imaged and analyzed with respect to the apical tissue surface. We found that the three-dimensional architecture of epithelial monolayers can vary widely even within small culture wells, and that layers that appear organized in the plane of the tissue can show gross disorganization in the apical-basal plane. Epithelial cell shapes should be analyzed in 3D to understand the architecture and maturity of the cultured tissue to accurately compare between experiments. Here, we present a detailed protocol for the use of our image analysis pipeline, Automated Layer Analysis (ALAn), developed to quantitatively characterize the architecture of cultured epithelial layers. ALAn is based on a set of rules that are applied to the spatial distributions of DNA and actin signals in the apical-basal (depth) dimension of cultured layers obtained from imaging cultured cell layers using a confocal microscope. ALAn facilitates reproducibility across experiments, investigations, and labs, providing users with quantitative, unbiased characterization of epithelial architecture and maturity.

Key features

• This protocol was developed to spatially analyze epithelial monolayers in an automated and unbiased fashion.

• ALAn requires two inputs: the spatial distributions of nuclei and actin in cultured cells obtained using confocal fluorescence microscopy.

• ALAn code is written in Python3 using the Jupyter Notebook interactive format.

• Optimized for use in Marbin-Darby Canine Kidney (MDCK) cells and successfully applied to characterize human MCF-7 mammary gland–derived and Caco-2 colon carcinoma cells.

• This protocol utilizes Imaris software to segment nuclei but may be adapted for an alternative method. ALAn requires the centroid coordinates and volume of nuclei.

Graphical overview

The advent of single-cell RNA sequencing (scRNAseq) has enabled in-depth gene expression analysis of several thousand cells isolated from tissues. We recently reported the application of scRNAseq toward the dissection of the tumor-infiltrating T-cell repertoire in human pancreatic cancer samples. In this study, we demonstrated that combined whole transcriptome and T-cell receptor (TCR) sequencing provides an effective way to identify tumor-reactive TCR clonotypes on the basis of gene expression signatures. An important aspect in this respect was the experimental validation of TCR-mediated anti-tumor reactivity by means of an in vitro functional assay, which is the subject of the present protocol. This assay involves the transient transfection of mRNA gene constructs encoding TCRα/β pairs into a well-defined human T-cell line, followed by co-cultivation with the tumor cells of interest and detection of T-cell activation by flow cytometry. Due to the high transfectability and the low background reactivity of the mock-transfected T-cell line to a wide variety of tumor cells, this assay offers a highly robust and versatile platform for the functional screening of large numbers of TCR clonotypes as identified in scRNAseq data sets. Whereas the assay was initially developed to test TCRs of human origin, it was more recently also applied successfully for the screening of TCRs of murine origin.

In vivo brain imaging, using a combination of genetically encoded Ca²⁺ indicators and gradient refractive index (GRIN) lens, is a transformative technology that has become an increasingly potent research tool over the last decade. It allows direct visualisation of the dynamic cellular activity of deep brain neurons and glia in conscious animals and avoids the effect of anaesthesia on the network. This technique provides a step change in brain imaging where fibre photometry combines the whole ensemble of cellular activity, and multiphoton microscopy is limited to imaging superficial brain structures either under anaesthesia or in head-restrained conditions. We have refined the intravital imaging technique to image deep brain nuclei in the ventral medulla oblongata, one of the most difficult brain structures to image due to the movement of brainstem structures outside the cranial cavity during free behaviour (head and neck movement), whose targeting requires GRIN lens insertion through the cerebellum—a key structure for balance and movement. Our protocol refines the implantation method of GRIN lenses, giving the best possible approach to image deep extracranial brainstem structures in awake rodents with improved cell rejection/acceptance criteria during analysis. We have recently reported this method for imaging the activity of retrotrapezoid nucleus and raphe neurons to outline their chemosensitive characteristics. This revised method paves the way to image challenging brainstem structures to investigate their role in complex behaviours such as breathing, circulation, sleep, digestion, and swallowing, and could be extended to image and study the role of cerebellum in balance, movement, motor learning, and beyond.

Key features

• We developed a protocol that allows imaging from brainstem neurons and glia in freely behaving rodents.

• Our refined method of GRIN lenses implantation and cell sorting approach gives the highest number of cells with the least postoperative complications.

• The revised deep brainstem imaging method paves way to understand complex behaviours such as cardiorespiratory regulation, sleep, swallowing, and digestion.

• Our protocol can be implemented to image cerebellar structures to understand their role in key functions such as balance, movement, motor learning, and more.

Graphical overview

Stem cell spheroids are rapidly becoming essential tools for a diverse array of applications ranging from tissue engineering to 3D cell models and fundamental biology. Given the increasing prominence of biotechnology, there is a pressing need to develop more accessible, efficient, and reproducible methods for producing these models. Various techniques such as hanging drop, rotating wall vessel, magnetic levitation, or microfluidics have been employed to generate spheroids. However, none of these methods facilitate the easy and efficient production of a large number of spheroids using a standard 6-well plate. Here, we present a novel method based on pellet culture (utilizing U-shaped microstructures) using a silicon mold produced through 3D printing, along with a detailed and illustrated manufacturing protocol. This technique enables the rapid production of reproducible and controlled spheroids (for 1 × 10⁶ cells, spheroids = 130 ± 10 μm) from human induced pluripotent stem cells (hIPSCs) within a short time frame (24 h). Importantly, the method allows the production of large quantities (2 × 10⁴ spheroids for 1 × 10⁶ cells) in an accessible and cost-effective manner, thanks to the use of a reusable mold. The protocols outlined herein are easily implementable, and all the necessary files for the method replication are freely available.

Key features

• Provision of 3D mold files (STL) to produce silicone induction device of spheroids using 3D printing.

• Cost-effective, reusable, and autoclavable device capable of generating up to 1.2× 10⁴ spheroids of tunable diameters in a 6-well plate.

• Spheroids induction with multiple hIPSC cell lines.

• Robust and reproducible production method suitable for routine laboratory use.

Graphical overview

Spheroid induction process following the pellet method on molded silicon discs

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