Cancer Biology

Macropinocytosis is an evolutionarily conserved process, which is characterized by the formation of membrane ruffles and the uptake of extracellular fluid. We recently demonstrated a role for CYFIP-related Rac1 Interactor (CYRI) proteins in macropinocytosis. High-molecular weight dextran (70kDa or higher) has generally been used as a marker for macropinocytosis because it is too large to fit in smaller endocytic vesicles, such as those of clathrin or caveolin-mediated endocytosis. Through the use of an image-based dextran uptake assay, we showed that cells lacking CYRI proteins internalise less dextran compared to their wild-type counterparts. Here, we will describe a step-by-step experimentation procedure to detect internalised dextran in cultured cells, and an image pipeline to analyse the acquired images, using the open-access software ImageJ/Fiji. This protocol is detailed yet simple and easily adaptable to different treatment conditions, and the analysis can also be automated for improved processing speed.

Cell migration is a vital process in the development of multicellular organisms. When deregulated, it is involved in many diseases such as inflammation and cancer metastisation. Some cancer cells could be stimulated using chemoattractant molecules, such as growth factor Heregulin β1. They respond to the attractant or repellent gradients through a process known as chemotaxis. Indeed, chemotactic cell motility is crucial in tumour cell dissemination and invasion of distant organs. Due to the complexity of this phenomenon, the majority of available in vitro methods to study the chemotactic motility process have limitations and are mainly based on endpoint assays, such as the Boyden chamber assay. Nevertheless, in vitro time-lapse microscopy represents an interesting opportunity to study cell motility in a chemoattracting gradient, since it generates large volume image-based information, allowing the analysis of cancer cell behaviours. Here, we describe a detailed time-lapse imaging protocol, designed for tracking T47D human breast cancer cell line motility, toward a gradient of Heregulin β1 in a Dunn chemotaxis chamber assay. The protocol described here is readily adapted to study the motility of any adherent cell line, under various conditions of chemoattractant gradients and of pharmacological drug treatments. Moreover, this protocol could be suitable to study changes in cell morphology, and in cell polarity.

Research on cell migration and interactions with the extracellular matrix (ECM) was mostly focused on 2D surfaces in the past. Many recent studies have highlighted differences in migratory behaviour of cells on 2D surfaces compared to complex cell migration modes in 3D environments. When embedded in 3D matrices, cells constantly sense the physicochemical, topological and mechanical properties of the ECM and adjust their behaviour accordingly. Changes in the stiffness of the ECM can have effects on cell morphology, differentiation and behaviour and cells can follow stiffness gradients in a process called durotaxis. Here we introduce a detailed protocol for the assembly of 3D matrices consisting of collagen I/fibronectin and embedding cells for live cell imaging. Further, we will show how the matrix can be stiffened via non-enzymatic glycation and how collagen staining with fluorescent dyes allows simultaneous imaging of both matrix and cells. This approach can be used to image cell migration in 3D microenvironments with varying stiffness, define cell-matrix interactions and the cellular response to changing ECM, and visualize matrix deformation by the cells.

Cell migration is a fundamental cellular process that plays a crucial role in many physioglogical and pathological processes such as wound healing or cancer metastasis. Many assays have been developed to examine cell migration, such as the wound healing or scratch assay, Boyden Chamber or transwell assay, and the method we will describe here, single cell migration assay. In this assay, cells are plated sparsely on a collagen coated plate and live cell imaging is performed over a period of 2 h at 1 frame per minute. After imaging is completed, cells are tracked manually using ImageJ by tracking movement of the centroid of the cell. These data points are then exported and overall distance travelled from frame to frame is determined and divided by total time imaged to determine speed of the cell. This method provides a quick way to examine effect of cellular manipulation on cell migration before proceeding to perform more complex assays.

The ability of cancer cells to migrate through a complex three-dimensional (3D) environment is a hallmark event of cancer metastasis. Therefore, an in vitro migration assay to evaluate cancer cell migration in a 3D setting is valuable to examine cancer progression. Here, we describe such a simple migration assay in a 3D collagen-fibronectin gel for observing cell morphology and comparing the migration abilities of cancer cells. We describe below how to prepare the collagen-fibronectin gel castings, how to set up time-lapse recording, how to draw single-cell trajectories from movies and extract key parameters that characterize cell motility, such as cell speed, directionality, mean square displacement, and directional persistence. In our set-up, cells are sandwiched in a single plane between two collagen-fibronectin gels. This trick facilitates the analysis of cell tracks, which are for the most part 2D, at least in the beginning, but in a 3D environment. This protocol has been previously published in Visweshwaran et al. (2018) and is described here in more detail.

The high migration rate of tumor cells often results in poor prognosis for the survival of the patients. Here, we describe a protocol to measure the migration of cells using a quantitative assay. The relative tumor cell migration was measured using ThinCerts^TM cell culture inserts and a lactate dehydrogenase (LDH) assay to quantify the relative cell number. The quantification of the migration with the LDH kit is much more precise than other methods using i.e. crystal blue to count the cells.

Cell migration is a highly complex and dynamic biological process, essential in several physiological phenomena and pathologies including cancer dissemination and metastasis formation. Thus understanding single cell migration is highly relevant and requires a suitable image-based assay. Depending on the speed of the moving cells, one may require fast time-lapse microscopy, which is not always suitable for high-throughput screening. To overcome this, a quantitative and fixed single cell migration assay was developed based on the PhagoKinetic Tracks (PKT) procedure. Briefly, cells are seeded on top of a monolayer of carboxylated latex beads, and as cells migrate, they phagocytose these beads and leave behind a migratory track. These bead-free migratory tracks can be visualized using a standard bright field microscope and analysed for a multiparametric quantitative assessment of single cell migration (Naffar-Abu-Amara et al., 2008).

Here we describe a detailed and optimized protocol of the PKT assay, adaptable for both RNAi and drug screening (van Roosmalen et al., 2015). This protocol allows the user to study migratory behaviour at the single cell level, without fast and live-imaging microscopy.

The scratch wound healing assay has been widely adapted and modified to study the effects of a variety of experimental conditions, for instance, gene knockdown or chemical exposure, on mammalian cell migration and proliferation. In a typical scratch wound healing assay, a “wound gap” in a cell monolayer is created by scratching, and the “healing” of this gap by cell migration and growth towards the center of the gap is monitored and often quantitated. Factors that alter the motility and/or growth of the cells can lead to increased or decreased rate of “healing” of the gap (Lampugnani, 1999). This assay is simple, inexpensive, and experimental conditions can be easily adjusted for different purposes. The assay can also be used for a high-throughput screen platform if an automated system is used (Yarrow and Perlman, 2004).