Mesenchymal stromal/stem cells (MSCs) are promising candidates for cell therapy and regenerative medicine. However, murine bone marrow-derived MSCs (mBMSCs) are much more difficult to isolate and purify because of their high heterogeneity and very limited numbers within bone morrow. To this end, this protocol was developed to establish an appropriate method for isolation and expansion of mBMSCs for preclinical research. Cells were dissociated and isolated mainly from compact bones via an enzyme digestion method to dissociate sufficient cell pools for further expansion ex vivo. Adherent cells collected were spindle-shaped with colony-forming units, and expressed a series of surface markers, including Oct4, CD29, and Sca-1 (stem cell antigen-1). Additionally, mBMSCs possessed potent capacity to differentiate into osteoblasts, adipocytes, and chondrocytes under appropriate culture conditions. Therefore, this study provides a standardized and reliable method for quick expansion of murine BMSCs. © 2026 Wiley Periodicals LLC. Basic Protocol: Enzymatic digestion method.
Biomaterial-based bioinks are increasingly utilized in bioprinting to engineer three-dimensional (3D) constructs with living cells for tissue engineering and disease modeling. Among various bioinks explored, alginate-based formulations stand out due to their good biocompatibility, mild gelation conditions, tunable mechanical properties, and ease of crosslinking via divalent cations such as calcium. Despite their widespread use, standardized protocols for preparing alginate-based bioinks and characterizing bioprinted constructs have not been well documented. Our laboratory has developed and validated reproducible methods for preparing a variety of alginate-based bioinks and printing cell-laden constructs tailored for diverse applications. In this article, we present detailed step-by-step protocols covering bioink preparation and rheological characterization, extrusion-based bioprinting of cell-laden constructs, post-printing culture and co-culture techniques, printability assessment, and live/dead and immunofluorescence assays. These protocols serve as a standardized framework for the fabrication and characterization of 3D bioprinted alginate-based cell-laden constructs, thereby facilitating translational research in tissue engineering, disease modeling, and preclinical therapeutic development. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1 : Bioink preparation Basic Protocol 2 : Bioink characterization using rheology Basic Protocol 3 : Scaffold design and bioprinting Support Protocol : 3D-printing parameter determination Basic Protocol 4 : Printability and cell viability analyses, and immunofluorescence assay
Angiogenesis is crucial in tissue repair, wound healing, and embryo development. Maintaining a balance between pro-angiogenic and anti-angiogenic factors is crucial for tissue homeostasis, thereby preventing the development of pathological conditions. In addition, the tissue microenvironment constantly influences molecular signaling, potentially altering this delicate balance. The chorioallantoic membrane (CAM) assay has emerged as a dynamic, cost-effective, and ethically favorable alternative for studying angiogenesis. This work presents a detailed and accessible approach to the CAM assay, focusing on the evaluation of the angiogenic potential of applied stimuli (such as conditioned media, extracellular vesicles, or bioscaffolds) through various techniques, including hematoxylin and eosin, PicroSirius Red, Alcian Blue, and resorcin-fuchsin staining; immunofluorescence; immunohistochemistry; and scanning electron microscopy (SEM). Our protocol uses the in ovo CAM assay approach, utilizing a simple incubator setup that eliminates the need for expensive equipment. The primary objective is to provide a comprehensive methodology for researchers to efficiently analyze angiogenesis, offering insights into vessel morphology and protein expression. The experiment comprises three major steps: (1) egg opening, stimulation, and sample collection; (2) histological processing and paraffin blocking; and (3) microscopic analyses. The first and second procedures take approximately 15 days, after which each analysis can be conducted whenever the researcher judges appropriate. The expected results include several types of data concerning the proliferation rate, vessel presence and integrity, measurement of angiogenic-specific proteins, and investigation of structural proteins. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol: Angiogenesis quail and chicken chorioallantoic membrane assay analyses.
Measuring oxygen consumption allows for the role of mitochondrial function in biological phenomena and mitochondrial diseases to be determined. Although respirometry has become a common approach in disease research, current methods are limited by the necessity to process and measure tissue samples within 1 hr of acquisition. Detailed by Acin-Perez and colleagues, a new respirometry approach designed for previously frozen tissue samples eliminates these hurdles for mitochondrial study. This technique allows for the measurement of maximal respiratory capacity in samples frozen for long-term storage before testing. This protocol article describes the optimal tissue isolation methods and the combination of substrates to define electron transport chain function at high resolution in previously frozen tissue samples. © 2020 The Authors. Basic Protocol 1: Sample collection, storage, and homogenization for previously frozen tissue respirometry Basic Protocol 2: Running a Seahorse respirometry assay using previously frozen tissue samples Basic Protocol 3: Normalization to mitochondrial content for previously frozen tissue respirometry.
Incorporation of a stable-isotope metabolic tracer into cells or tissue can be followed at submicron resolution by multi-isotope imaging mass spectrometry (MIMS), a form of imaging secondary ion microscopy optimized for accurate isotope ratio measurement from microvolumes of sample (as small as ∼30 nm across). In a metabolic MIMS experiment, a cell or animal is metabolically labeled with a tracer containing a stable isotope. Relative accumulation of the heavy isotope in the fixed sample is then measured as an increase over its natural abundance by MIMS. MIMS has been used to measure protein turnover in single organelles, track cellular division in vivo, visualize sphingolipid rafts on the plasma membrane, and measure dopamine incorporation into dense-core vesicles, among other biological applications. In this article, we introduce metabolic analysis using NanoSIMS by focusing on two specific applications: quantifying protein turnover in single organelles of cultured cells and tracking cell replication in mouse tissues in vivo. These examples illustrate the versatility of metabolic analysis with MIMS. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Metabolic labeling for MIMS Basic Protocol 2: Embedding of samples for correlative transmission electron microscopy and MIMS with a genetically encoded reporter Alternate Protocol: Embedding of samples for correlative light microscopy and MIMS Support Protocol: Preparation of silicon wafers as sample supports for MIMS Basic Protocol 3: Analysis of MIMS data.
Eukaryotic integral membrane proteins are key components of various biological processes. Because they are implicated in multiple diseases, it is important to understand their mechanism of action by elucidating their structure and function. Complex technical challenges associated with the generation of recombinant membrane proteins severely impair our ability to understand them using structural and biochemical methods. Here, we provide a detailed procedure to address and mitigate difficulties involved in the large-scale heterologous overexpression and purification of eukaryotic membrane proteins using HEK293S GnTi- cells transduced with baculovirus. Two human proteins, hDHHC15 and hPORCN, are presented as examples, with step-by-step instructions for transient transfection and generation of baculoviruses, followed by overexpression and purification from HEK293S GnTi- cells. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Small-scale protein expression in mammalian HEK293T cells Basic Protocol 2: Generation of baculovirus from Sf9 (insect) cells Alternate Protocol: Enumeration-free method for generating P2 viral stock Support Protocol 1: Small-scale transduction of HEK293T cells with P2 baculovirus Basic Protocol 3: Large-scale viral transduction of HEK293S GnTi- cells Support Protocol 2: Large-scale membrane preparation from HEK293S GnTi- cells Basic Protocol 4: Large-scale purification of membrane proteins from HEK293S GnTi- cells.
Phagocytes, notably macrophages, are critical sentinels of their environment, patrolling for and eradicating unwanted components. The ability of cells to process extracellular cargo in an appropriate manner is important for both clearance of the cargo and eventual return to homeostasis. Although the evolutionarily conserved pathway of autophagy involves the degradation and recycling of unnecessary or dysfunctional cellular components during starvation, we now appreciate that the reach of autophagy extends beyond nutrient deprivation, notably including cellular quality control (e.g., mitophagy) and host defense against internalized pathogens (i.e., xenophagy). Despite being seemingly disparate, autophagic functions are unified as conserved mechanisms for containment and immunosuppression, suggesting an original immune function for autophagy. A recently described pathway called LC3-associated phagocytosis (LAP) marries the ancient concepts of phagocytosis and autophagy, revealing new ways in which the autophagy machinery, in a molecularly distinct pathway, contributes to the inflammatory response. In this article, protocols to detect LAP by electron microscopy, immunofluorescence, flow cytometry, and phagosome purification are described, allowing the user to detect multiple characteristics of LAP in both qualitative and quantitative manners. Published 2020. U.S. Government. Basic Protocol 1: Detection of LAP by electron microscopy Basic Protocol 2: Detection of LAP by confocal microscopy of LC3-GFP-expressing cells Alternate Protocol 1: Detection of LAP by confocal microscopy using immunofluorescence Basic Protocol 3: Detection of LAP using flow cytometry of LC3-GFP-expressing cells Alternate Protocol 2: Detection of LAP using antibody staining and flow cytometry Basic Protocol 4: Detection of LAP by western blot of purified LAPosomes.
The movement of intracellular cargo, such as transcripts, proteins, and organelles, is fundamental to cellular function. Neurons, due to their long axons and dendrites, are particularly dependent on proper intracellular trafficking and vulnerable to defects in the movement of intracellular cargo that are noted in neurodegenerative and neurodevelopmental disorders. Accurate quantification of intracellular transport is therefore needed for studying the mechanisms of cargo trafficking, the influence of mutations, and the effects of potentially therapeutic pharmaceuticals. In this article, we introduce an algorithm called "Kymolyzer." The algorithm can quantify intracellular trafficking along a defined path, such as that formed by the aligned microtubules of axons and dendrites. Kymolyzer works as a semi-autonomous kymography software application. It constructs and analyzes kymographs to measure the movement and distribution of fluorescently tagged objects along a user-defined path. The algorithm can be used under a wide variety of experimental conditions and can extract a diverse array of motility parameters describing intracellular movement, including time spent in motion, percentage of objects in motion, percentage of objects that are stationary, and velocities of motile objects. This article serves as a user manual describing the design of Kymolyzer, providing a stepwise protocol for its use and illustrating its functions with common examples. © 2020 Wiley Periodicals LLC Basic Protocol: Kymolyzer, a semi-autonomous kymography tool to analyze intracellular motility.
Spatiotemporal localization of protein function is essential for physiological processes from subcellular to tissue scales. Genetic and pharmacological approaches have played instrumental roles in isolating molecular components necessary for subcellular machinery. However, these approaches have limited capabilities to reveal the nature of the spatiotemporal regulation of subcellular machineries like those of cytoskeletal organelles. With the recent advancement of optogenetic probes, the field now has a powerful tool to localize cytoskeletal stimuli in both space and time. Here, we detail the use of tunable light-controlled interacting protein tags (TULIPs) to manipulate RhoA signaling in vivo. This is an optogenetic dimerization system that rapidly, reversibly, and efficiently directs a cytoplasmic RhoGEF to the plasma membrane for activation of RhoA using light. We first compare this probe to other available optogenetic systems and outline the engineering logic for the chosen recruitable RhoGEFs. We also describe how to generate the cell line, spatially control illumination, confirm optogenetic control of RhoA, and mechanically induce cell-cell junction deformation in cultured tissues. Together, these protocols detail how to probe the mechanochemical circuitry downstream of RhoA signaling. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Generation of a stable cell line expressing TULIP constructs Basic Protocol 2: Preparation of collagen substrate for imaging Basic Protocol 3: Transient transfection for visualization of downstream effectors Basic Protocol 4: Calibration of spatial illumination Basic Protocol 5: Optogenetic activation of a region of interest.
Mitophagy is a selective autophagy process that specifically removes damaged mitochondria via general autophagy. The two major recessive Parkinson's disease genes PINK1 and Parkin play essential roles in mitophagy initiation, increasing the interest in mitophagy in both basic and translational research over the past 10 years. Initially, mitophagy was measured by the loss of mitochondria either through confocal imaging or immunoblot of mitochondrial proteins such as Tom20 or COXII. Confocal imaging of mitochondria DNA loss via anti-DNA staining is another option. All of these methods, which take considerable effort and labor, are not sensitive enough to detect early stages of mitophagy with unambiguous objectivity. The mKeima assay can be used for both confocal imaging and FACS analysis to provide a thorough picture of mitophagy with a wide dynamic range. The Keima fluorophore has bimodal excitation under neutral and acidic pH conditions. Thus, when Keima is targeted to mitochondria it can accurately reveal the formation of mitolysosomes. Here the author briefly describes the origin and history of mKeima and how it is adapted to measure mitophagy. The author presents detailed protocols for making stable cell lines for optimized mitophagy detection and discuss many parameters that might affect the assay. A troubleshooting section is also provided to discuss possible pitfalls to improve reproducibility and sensitivity of the assay. © 2019 The Authors. Basic Protocol 1: Making stable lines expressing mito-mKeima and YFP-Parkin Support Protocol: Retrovirus/lentivirus infection Basic Protocol 2: FACS analysis of mitophagy.
Biochemical methods can reveal stable protein-protein interactions occurring within cells, but the ability to observe transient events and to visualize the subcellular localization of protein-protein interactions in cells and tissues in situ provides important additional information. The Proximity Ligation Assay® (PLA) offers the opportunity to visualize the subcellular location of such interactions at endogenous protein levels, provided that the probes that recognize the target proteins are within 40 nm. This sensitive technique not only elucidates protein-protein interactions, but also can reveal post-translational protein modifications. The technique is useful even in cases where material is limited, such as when paraffin-embedded clinical specimens are the only available material, as well as after experimental intervention in 2D and 3D model systems. Here we describe the basic protocol for using the commercially available Proximity Ligation Assay™ materials (Sigma-Aldrich, St. Louis, MO), and incorporate details to aid the researcher in successfully performing the experiments. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Proximity ligation assay Support Protocol 1: Antigen retrieval method for formalin-fixed, paraffin-embedded tissues Support Protocol 2: Creation of custom PLA probes using the Duolink™ In Situ Probemaker Kit when commercially available probes are not suitable Basic Protocol 2: Imaging, quantification, and analysis of PLA signals.
Science and medicine have become increasingly "human-centric" over the years. A growing shift away from the use of animals in basic research has led to the development of sophisticated in vitro models of various tissues utilizing human-derived cells to study physiology and disease. The human cornea has likewise been modeled in vitro using primary cells derived from corneas obtained from cadavers or post-transplantation. By utilizing a cell's intrinsic ability to maintain its tissue phenotype in a pre-designed microenvironment containing the required growth factors, physiological temperature, and humidity, tissue-engineered corneas can be grown and maintained in culture for relatively long periods of time on the scale of weeks to months. Due to its transparency and avascularity, the cornea is an optimal tissue for studies of extracellular matrix and cell-cell interactions, toxicology and permeability of drugs, and underlying mechanisms of scarring and tissue regeneration. This paper describes methods for the cultivation of corneal keratocytes, fibroblasts, epithelial, and endothelial cells for in vitro applications. We also provide detailed, step-by-step protocols for assembling and culturing 3D constructs of the corneal stroma, epithelial- and endothelial-stromal co-cultures and isolation of extracellular vesicles. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Isolating and culturing human corneal keratocytes and fibroblasts Basic Protocol 2: Isolating and culturing human corneal epithelial cells Basic Protocol 3: Isolating and culturing human corneal endothelial cells Basic Protocol 4: 3D corneal stromal construct assembly Basic Protocol 5: 3D corneal epithelial-stromal construct assembly Basic Protocol 6: 3D corneal endothelial-stromal construct assembly Basic Protocol 7: Isolating extracellular vesicles from corneal cell conditioned medium Support Protocol: Cryopreserving human corneal fibroblasts, corneal epithelial cells, and corneal endothelial cells.
BioID, a proximity biotinylation technique, offers a valuable approach to examine the interactions occurring within protein complexes that complements traditional protein biochemical methods. BioID has various advantages that are beneficial to the study of complexes, including an ability to detect insoluble and transient proteins. We have applied BioID to the study of integrin adhesion complexes (IACs), which are located at the junction between the plasma membrane and actin cytoskeleton. The use of multiple BioID baits enables a complex-wide, spatial annotation of IACs, which in turn facilitates the detection of novel proximal interactors and provides insights into IAC architecture. This article describes the labeling and affinity purification of IAC-proximal proteins and their analysis by label-free quantitative mass spectrometry. The article also outlines steps to identify high-confidence proximity interactors, and to interrogate the topology and functional relevance of proximity interaction networks through bioinformatic analyses. © 2020 The Authors. Basic Protocol 1: Proximity biotinylation of integrin adhesion complex components Basic Protocol 2: Mass spectrometry data processing by MaxQuant and detection of high-confidence proximal interactors Basic Protocol 3: Bioinformatic analysis and data visualization.
Construction of organized three-dimensional (3D) tissue with extracellular matrix (ECM) and multiple types of cells is important for tissue engineering to enable tissue function and enhance cellular function. However, the concentration of ECM and the thickness of the 3D tissue have been limited in previous methods due to a lack of permeability to nutrients and oxygen. Besides, it is difficult to use matured natural ECM as a cell scaffold without chemical modification due to its insolubility. In this article, we focus on multi-layered structure, which is commonly found in living tissue such as skin, blood vessels, and other organs. Here, we describe the preparation of a paper-like scaffold (ECM paper) from micro-fibered natural ECM and the construction of 3D multi-layered tissue composed of cell layers and ECM layers by stacking cell-seeded ECM papers. The thickness and components of the ECM layers are easily controllable by changing the composition of the ECM papers, and the fibrous structure of ECM paper shows high permeability and permits cell migration. Additionally, the ECM microfiber, which is physically defiberized from natural ECM, has a high ECM concentration equal to that of living tissue and high stability under physiological conditions. Therefore, this set of protocols enables construction of multi-layered 3D tissue composed of precisely controlled ECM layers and cell layers that may contribute to the assembly of tissue models. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Preparation of extracellular matrix paper Basic Protocol 2: Evaluation of cellular function of cells on extracellular matrix paper Basic Protocol 3: Construction of multi-layered 3D tissue.
Exosomes are 50- to 150-nm-diameter extracellular vesicles secreted by all mammalian cells except mature red blood cells and contribute to diverse physiological and pathological functions within the body. Many methods have been used to isolate and analyze exosomes, resulting in inconsistencies across experiments and raising questions about how to compare results obtained using different approaches. Questions have also been raised regarding the purity of the various preparations with regard to the sizes and types of vesicles and to the presence of lipoproteins. Thus, investigators often find it challenging to identify the optimal exosome isolation protocol for their experimental needs. Our laboratories have compared ultracentrifugation and commercial precipitation- and column-based exosome isolation kits for exosome preparation. Here, we present protocols for exosome isolation using two of the most commonly used methods, ultracentrifugation and precipitation, followed by downstream analyses. We use NanoSight nanoparticle tracking analysis and flow cytometry (Cytek® ) to determine exosome concentrations and sizes. Imaging flow cytometry can be utilized to both size exosomes and immunophenotype surface markers on exosomes (ImageStream® ). High-performance liquid chromatography followed by nano-flow liquid chromatography-mass spectrometry (LCMS) of the exosome fractions can be used to determine the presence of lipoproteins, with LCMS able to provide a proteomic profile of the exosome preparations. We found that the precipitation method was six times faster and resulted in a ∼2.5-fold higher concentration of exosomes per milliliter compared to ultracentrifugation. Both methods yielded extracellular vesicles in the size range of exosomes, and both preparations included apoproteins. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Pre-analytic fluid collection and processing Basic Protocol 2: Exosome isolation by ultracentrifugation Alternate Protocol 1: Exosome isolation by precipitation Basic Protocol 3: Analysis of exosomes by NanoSight nanoparticle tracking analysis Alternate Protocol 2: Analysis of exosomes by flow cytometry and imaging flow cytometry Basic Protocol 4: Downstream analysis of exosomes using high-performance liquid chromatography Basic Protocol 5: Downstream analysis of the exosome proteome using nano-flow liquid chromatography-mass spectrometry.
Neural crest cells constitute a unique population of progenitor cells with extensive stem cell capacities able to navigate throughout various environments in the embryo and are a source of multiple cell types, including neurons, glia, melanocytes, smooth muscles, endocrine cells, cardiac cells, and also skeletal and supportive tissues in the head. Neural crest cells are not restricted to the embryo but persist as well in adult tissues where they provide a reservoir of stem cells with great therapeutic promise. Many fundamental questions in cell, developmental, and stem cell biology can be addressed using this system. During the last decades there has been an increased availability of elaborated techniques, animal models, and molecular tools to tackle neural crest cell development. However, these approaches are often very challenging and difficult to establish and they are not adapted for rapid functional investigations of mechanisms driving cell migration and differentiation. In addition, they are not adequate for collecting pure populations of neural crest cells usable in large scale analyses and for stem cell studies. Transferring and adapting the neural crest system in tissue culture may then represent an attractive alternative, opening up numerous prospects. Here we describe a simple method for establishing primary cultures of neural crest cells derived from trunk neural tubes using the avian embryo as a source of cells. This protocol is suited for producing pure populations of neural crest cells that can be processed for cytological, cellular, and functional approaches aimed at characterizing their phenotype, behavior, and potential. © 2020 Wiley Periodicals LLC. Basic Protocol : Primary cultures of avian trunk neural crest cells Support Protocol : Adaptations for immunofluorescence labeling and videomicroscopy
Trafficking of intracellular cargo is essential to cellular function and can be defective in pathological states including cancer and neurodegeneration. Tools to quantify intracellular traffic are thus necessary for understanding this fundamental cellular process, studying disease mechanisms, and testing the effects of therapeutic pharmaceuticals. In this article we introduce an algorithm called QuoVadoPro that autonomously quantifies the movement of fluorescently tagged intracellular cargo. QuoVadoPro infers the extent of intracellular motility based on the variance of pixel illumination in a series of time-lapse images. The algorithm is an unconventional approach to the automatic measurement of intracellular traffic and is suitable for quantifying movements of intracellular cargo under diverse experimental paradigms. QuoVadoPro is particularly useful to measure intracellular cargo movement in non-neuronal cells, where cargo trafficking occurs as short movements in mixed directions. The algorithm can be applied to images with low temporal or spatial resolutions and to intracellular cargo with varying shapes or sizes, like mitochondria or endoplasmic reticulum: situations in which conventional methods such as kymography and particle tracking cannot be applied. In this article we present a stepwise protocol for using the QuoVadoPro software, illustrate its methodology with common examples, discuss critical parameters for reliable data analysis, and demonstrate its use with a previously published example. © 2020 Wiley Periodicals LLC. Basic Protocol: QuoVadoPro, an autonomous tool for measuring intracellular dynamics using temporal variance.
In the past, in vitro studies of invasion and tumor progression were performed primarily using cancer cells cultured on a flat, two-dimensional (2D) surface in a monolayer. In recent years, however, many studies have demonstrated differences in cell signaling and cell migration between 2D and 3D cell cultures. Traditional 2D monolayer cancer cell invasion models do not fully recapitulate 3D cell-to-cell and cell-to-extracellular matrix interactions that in vivo models can provide. Moreover, although in vivo animal models are irreplaceable for studying tumor biology and metastasis, they are costly, time-consuming, and impractical for answering preliminary questions. Thus, emergent and evolving 3D spheroid cell culture models have changed the way we study tumors and their interactions with their surrounding extracellular matrix. In the case of breast cancer, metastasis of breast cancer tumors results in high mortality rates, and thus development of robust cell culture models that are reproducible and practical for studying breast cancer progression is important for ultimately developing preventatives for cancer metastasis. This article provides a set of protocols for generating uniform spheroids with a thin sheet of basement membrane for studying the initial invasion of mammary epithelial cells into a surrounding collagen-rich extracellular matrix. Details are provided for generating 3D spheroids with a basement membrane, polymerizing collagen I, embedding the spheroids in the 3D collagen gel, and immunostaining the spheroids for invasion studies. Published 2020. U.S. Government. Basic Protocol 1: Growth of uniformly sized tumor spheroids with an encapsulating basement membrane Basic Protocol 2: Polymerization and embedding of tumor spheroids in a 3D type I collagen gel Alternate Protocol: Embedding of tumor spheroids in collagen gels using a sandwich method Basic Protocol 3: Fixing and immunostaining of tumor spheroids embedded in 3D collagen gels.
We developed a modified invasion assay in three-dimensional (3D) gels that permits isolation of invading cells as living cells, termed an invading cell-trapping (iCT) assay. A small cell strainer consisting of nylon mesh with pores of 40-µm square is used in this assay. A layer of gel composed of extracellular-matrix components is formed on each side of the nylon mesh, which permits cell migration or invasion from one gel layer to the other. At the end of the assay, the two gel layers are removed from the apparatus and easily separated from each other. Invading cells from the primary gel are trapped in the secondary gel, which maintains the morphology and other properties of the invasive cells in a 3D matrix. The cells that have invaded are observed and counted with a standard light microscope without cell staining. There is no need for a specialized microscope, imaging analysis software, or advanced cell-biological technical knowledge in this assay. This assay can also reduce measurement of nonspecific cell invasion by monitoring the upward invasion of cells. The viability of both invading and non-invading cells trapped in the gels can be assessed by typical colorimetric assays, if desired. This assessment characterizes the total number of cells (invading and non-invading cells) and the ratio of invading cells to total cells. By repeating the iCT assay, further enrichment of invasive and noninvasive cells can be attained. Thus, this assay improves comparative analyses between invasive and noninvasive cells. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Measuring upward cell invasion into collagen gel Basic Protocol 2: Measuring cell invasion from Matrigel into collagen gel Basic Protocol 3: Isolation and enrichment of highly invasive cells.
The explosive growth of Machine Learning provided scientists with insights into the data in the ways unattainable using established research techniques. It allowed the detection of biological features that were previously unrecognized and overlooked. Yet, since Machine Learning methodology originates from informatics, many cell biology laboratories experience difficulties with implementing it. In preparing this article, we targeted the rapidly expanding audience of cell and molecular biologists who perform analysis of microscopy images and seek to add Machine Learning models to their research workflow. We review the advantages of using Machine Learning in microscopy projects, describe the Machine Learning pipeline, and share practical guidelines for building the models. The latest developments in the rapidly expanding field are also given. The technical survey is concluded with an overview of the tools required for model creation and advice on their use. © 2023 Wiley Periodicals LLC.
