Name: measuring cell-edge protrusion dynamics during spreading using live-cell microscopy
Uploaded: 2021-11-01T13:00:00.000+00:00
Duration: 5 min 50 s
Description: Watch this Scientific Journal Video about Measuring Cell-Edge Protrusion Dynamics during Spreading using Live-Cell Microscopy at JoVE.com

This work was supported by grants NIH MH115939, NS112121, NS105640, and R56MH122449-01A1 (to Anthony J. Koleske) and from the Israel Science Foundation (grants number 1462/17 and 2142/21) (to Hava Gil-Henn).

All methods described in this protocol have been approved by the institutional Animal Care and Use Committee (IACUC) of Bar-Ilan University.
NOTE: A step-by-step graphical depiction of the procedure described in this section appears in Figure 1.
1. Cell culture
NOTE: The cells used in the protocol are mouse embryonic fibroblasts (MEFs) that were generated from E11.5-13.5 embryos of wild-type C57BL/6 mice. Primary MEFs were generated according to the Jacks laboratory protocol13. Cells from five different embryos were pooled together and immortalized by infection with a retroviral vector expressing SV40 large T antigen followed by selection with 4 mM Histidinol for 3 weeks.
<ol>
	<li>Culture cells in tissue culture plates containing Dulbecco&#39;s Modified Eagle Medium (DMEM) containing 1 g/L glucose, 1% glutamine, 1% penicillin-streptomycin, and 10% fetal bovine serum (FBS) (see Table of Materials), in a humidified incubator with 5% CO2 at 37 &#176;C.</li>
	<li>Culture the cells until 90%-95% confluency and split at a ratio of 1:5 every 2-3 days.</li>
</ol>
2. Glass-bottom dish coating
NOTE: Glass-bottom dish coating should be performed in the tissue culture hood in sterile conditions.
<ol>
	<li>Add 2 mL of 1N HCl solution at the center of a glass-bottom dish (Table of Materials) and incubate for 20 min at room temperature (RT). 
	NOTE: This stage is meant to remove residue from the glass that may interrupt imaging later.</li>
	<li>Wash the glass-bottom dish three times with 2 mL of 1x PBS (Table of Materials) each.</li>
	<li>Dilute fibronectin (see Table of Materials) to 10 &#181;g/mL in 1x PBS. Add 200 &#181;L of the diluted fibronectin solution to the glass center of the dish. Incubate at 37 &#176;C (tissue culture incubator) for 1 h. 
	NOTE: Alternatively, the glass-bottom dish can be incubated overnight at 4 &#176;C on a flat surface.</li>
	<li>During fibronectin coating incubation time, prepare 1% BSA (Table of Materials) solution in 1x PBS, filter through 0.2 &#181;m, and denature by incubating at 70 &#176;C for 30 min in a pre-warmed water-bath.</li>
	<li>Wash the coated glass-bottom dish three times with 2 mL of 1x PBS each.</li>
	<li>Add 2 mL of denatured BSA solution to the glass center and incubate for 1 h at 37 &#176;C (tissue culture incubator). 
	NOTE: Alternatively, the glass-bottom dish can be incubated overnight at 4 &#176;C on a flat surface.</li>
	<li>Wash the glass-bottom dish 3 times with 2 mL of 1x PBS each. 
	&#8203;NOTE: Incubation of glass-bottom dishes with fibronectin should be performed for at least 1 h at 37 &#176;C to coat the surface properly. Shorter incubation time may not produce proper coating, and as a result, cell phenotype may not be related to integrin activation. The BSA layer is inert and will not affect protrusion dynamics and is meant to block free potential sites for integrin-independent cell adhesion. The BSA must be denatured before coating to prevent it from inducing cell apoptosis, which can influence cell-edge dynamics.</li>
</ol>
3. Preparation of cells for imaging
<ol>
	<li>Sixteen to eighteen hours before the experiment, plate the cells to reach 70%-80% confluence on the following day. For MEFs, plate 0.7 x 106 cells per 10 cm diameter tissue culture plate a day before performing the experiment. 
	NOTE: For a successful protrusion experiment, it is important that the cells will be used during their logarithmic growth phase. To achieve this, cells should reach 70%-80% confluence on the day of the experiment. A higher density of cells will result in a longer spreading time and/or impediment in attachment during the experiment.</li>
	<li>On the experiment day, add 2 mL of trypsin solution (Table of Materials) per 10 cm diameter tissue culture plate and incubate for 2-3 min in a tissue culture incubator until cells detach. Inactivate trypsin by adding 5 mL of the complete medium.</li>
	<li>Count the cells using a hemocytometer and plate 20,000 cells in 2 mL of the complete medium on a glass-bottom dish coated as above (section 2). 
	NOTE: The number of cells for plating depends on the size and type of cells. For larger or more spread cells, plate only 10,000 cells to have enough cells for imaging. It is important to pick single cells that do not touch as cell-to-cell contact can alter cell-intrinsic spreading behaviors.</li>
	<li>Incubate the glass-bottom dishes with plated cells in the tissue culture incubator for 15 min. 
	&#8203;NOTE: During this stage, cells adhere to the fibronectin substrate and spread over it. When measuring protrusions during cell spreading, cells should be allowed to spread for 15 min after plating and before imaging. Imaging can be performed within a time window between 15 min to ~1 h following plating, and in any case, before the cells start migrating.</li>
</ol>
4. Microscope setup and imaging
NOTE: Various live-cell microscopy systems are available. The system used here is a Leica AF6000 inverted microscope equipped with CO2 and heating units and is attached to an ORCA-Flash 4.0 V2 digital CMOS camera.
<ol>
	<li>Turn on the heating unit at least 1 h before imaging and set it at 37 &#176;C.</li>
	<li>Turn on the CO2 unit at least 10 min before imaging and set it at 5% CO2.</li>
	<li>Switch on the microscope, camera, and computer.</li>
	<li>Open the microscope acquisition software (see Table of Materials). Choose the folder to save captured images and type a file name. Save every movie as a new file.</li>
	<li>Set the magnification on a 40x dry lens, phase-contrast. Set the time interval on 5 s, total movie duration 10 min.</li>
	<li>After 15 min incubation of the plated cells, place the glass-bottom dish with adhered cells into the adapter and fix it. Insert the adapter with the dish into its slot in the microscope stage.</li>
	<li>Take off the dish cover and place the CO2 lid instead. Open the CO2 valve. 
	&#8203;NOTE: Make sure the CO2 cover is clean before placing it on top of the glass-bottom dish. A dirty cover will reduce the quality of movies. Wipe the underside of the CO2 lid with a lint-free wipe soaked with 70% ethanol to remove dust and dirt. Wipe a second time with a dry lint-free wipe.</li>
	<li>View the cells and find an appropriate cell for imaging. Make sure the cell is in focus and start movie acquisition.</li>
</ol>
5. Image analysis
NOTE: Image analysis is performed using ImageJ (Table of Materials) as following:
<ol>
	<li>Open the acquired movie in ImageJ.</li>
	<li>Using the Straight tool in the main toolbar, make eight lines of 20 arbitrary units each perpendicular to the protrusions, including the lamella and cell edge, in a radial arrangement every 45&#176;, as shown in Figure 2A.</li>
	<li>In the main toolbar, go to Image &#62; Stacks &#62; Reslice. This will yield a kymograph picture, which describes the movement of single points within the cell membrane (Figure 2B). This action should be performed for every line out of the eight lines separately.</li>
	<li>Extract and manually count from the respective kymograph images the number of protrusions, retractions, and ruffles in each of the eight regions in the cell, marked by the grid lines. These numbers represent the frequency of protrusions, retractions, and ruffles per 10 min (Figure 2C, D). 
	NOTE: Ruffles can be distinguished from other structures based on their dark appearance in phase-contrast microscopy and their centripetal movement, which starts at the cell edge and ends at the border of the cell body, which can be observed in the acquired movies. Of note, when quantifying protrusion, retraction, and ruffle frequency, movies should be observed as a control for quantification and especially for defining ruffles.</li>
	<li>Determine the protrusion persistence, distance, and velocity by kymography analysis. For each of the generated kymographs, the X-axis represents distance, and the Y-axis represents time.</li>
	<li>To measure protrusion distance, follow steps 5.6.1-5.6.2.
	<ol>
		<li>Draw a perpendicular line from the base of the protrusion to the highest peak of the protrusion. Press M in ImageJ to measure the length of the line in pixels.</li>
		<li>To convert the length from pixels to &#956;m, ensure that the pixel to &#956;m ratio is known. 
		NOTE: The &#956;m to pixel ratio is the physical length of a pixel on the CCD camera / total magnification. The pixel size is characteristic of each type of camera. For example, for the camera used in this study, the pixel size is 6.5 &#956;m x 6.5 &#956;m, the physical length is 6.5 &#956;m and the magnification we used is 40x. Therefore, the &#956;m to pixel ratio of our camera is 0.1625 &#956;m/pixel. In the analysis, for a line in the length of 30 pixels, the protrusion distance would be 30 pixels x 0.1625 &#956;m/pixel = 4.875 &#956;m (Figure 3).</li>
	</ol>
	</li>
	<li>To measure protrusion time (persistence; the amount of time a protrusion spends protruding before retracting), follow steps 5.7.1-5.7.2.
	<ol>
		<li>Draw a horizontal line from the beginning of the protrusion (left to right) to the region of the highest peak. Press M in ImageJ to measure the length of the line in pixels.</li>
		<li>To convert the length from pixels to minutes, calculate the pixel to min ratio. This value depends on the interval between images. 
		NOTE: In this example, the interval between images is 5 s, the min/pixel ratio is 0.0833, and the horizontal line length is 8 pixels. Therefore, the protrusion time is 8 pixels x 0.0833 min/pixel = 0.6664 min.</li>
	</ol>
	</li>
	<li>Measure and calculate the retraction time similarly to protrusion time for a line drawn horizontally at the base of the protrusion from where the peak of the protrusion is to its base on the right (line X2 in Figure 3).</li>
	<li>Calculate the protrusion velocity by dividing protrusion distance by protrusion time. Calculate the retraction velocity by dividing protrusion distance by retraction time. In this example, protrusion velocity is calculated as 4.875 &#956;m/0.6664 min = 7.315 &#956;m/min., and the retraction velocity is identical because the line representing time is at the same length. 
	NOTE: In comparing different cell types, i.e., cells expressing wild type and mutant constructs of the same protein, it is imperative to perform a blinded analysis, so no bias is introduced.</li>
</ol>

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The authors have no conflict of interests to disclose.

Cell-edge protrusion dynamics, comprised of protrusions, retractions, and ruffles, is both a prerequisite and a potential rate-limiting event in cell motility. Here we describe a fast and simple method for measuring the dynamics of cell-edge protrusions during spreading. This method enables short-time imaging, generates a significant amount of data, does not require fluorescent labeling of cells or expensive fluorescent microscopy equipment, and could be used as a preliminary method for testing cytoskeletal dynamics invo...

Cell migration is a critical process that controls the development and function of all living organisms. Cell migration occurs in both physiological conditions, such as embryogenesis, wound healing, and immune response, and in pathological conditions, such as cancer metastasis and autoimmune disease. Despite differences in cell types that take part in different migratory events, all cell motility events share similar molecular mechanisms, which have been conserved in evolution from protozoa to mammals, and involve common cytoskeletal control mechanisms that can sense the environment, respond to signals, and modulate cell behavior in response1.
An initial stage in cell migration can be the formation of highly dynamic protrusions at the leading edge of the cell. Behind the lamellipodium one can find the lamella, which couples actin to myosin II-mediated contractility and mediates adhesion to the underlying substrate. Lamellipodia are induced by extracellular stimuli such as growth factors, cytokines, and cell adhesion receptors and are driven by actin polymerization, which provides the physical force that pushes the plasma membrane forward2,3. Many signaling and structural proteins have been implicated in this; among them are Rho GTPases, which act coordinately with other signals to activate actin-regulating proteins such as the Arp 2/3 complex, WASP family proteins, and members of the Formin and Spire families in lamellipodia2,4,5.
In addition to actin polymerization, myosin II activity is required for generating contractile forces at the lamellipodium and the anterior lamella. These contractions, also defined as cell-edge retractions, can also result from depolymerization of dendritic actin at the cell periphery and are critical for developing the lamellipodial leading edge and allowing the protrusion to sense the flexibility of the extracellular matrix and other cells and determine the direction of migration6,7,8. Cell edge protrusions that cannot attach to the substrate will form peripheral membrane ruffles, sheet-like structures that appear on the ventral surface of lamellipodia and lamella and move backward relative to the direction of migration. As the lamellipodium fails to attach to the substrate, a posterior lamellipodium forms underneath it and mechanically pushes the first lamellipodium toward the upper ventral surface. The actin filaments in the ruffle that were formerly parallel to the substrate now become perpendicular to it, and the ruffle is now positioned above the advancing lamellipodium. The ruffle that moves backward falls back into the cytosol and represents a cellular mechanism for recycling lamellipodial actin9,10.
Here, we describe an assay for the measurement of cell-edge protrusion dynamics. The protrusion assay uses time-lapse video microscopy to measure single cell-edge protrusion dynamics for 10 min during the spreading phase of the cell. Protrusion dynamics are analyzed by generating kymographs from these movies. In principle, a kymograph imparts detailed quantitative data of moving particles in a spatiotemporal plot to yield a qualitative understanding of cell edge dynamics. The intensity of the moving particle is plotted for all image stacks in a time versus space plot, where the X-axis and the Y-axis represent time and distance, respectively11. This method uses a manual kymograph analysis with ImageJ to get detailed quantitative data, enabling retrieving information from movies and images in case of low signal-to-noise ratio and/or high feature density, and the analysis of images acquired in phase-contrast light microscopy or poor image quality.
The cell-edge protrusion dynamics assay described herein is a fast, simple, and cost-effective method. As such, and because it has been shown to directly correlate with cell migration11,12, it can be used as a preliminary method for testing cytoskeletal dynamics involved in cell motility before deciding to perform more resource-demanding methods. Moreover, it also enables quantitative measurement of how genetic manipulations (knockout, knockdown, or rescue constructs) of cytoskeletal proteins impact cytoskeletal dynamics using a straightforward platform. The assay is an instructive model for exploring cytoskeletal dynamics in the context of cell migration and could be used for elucidation of the mechanisms and molecules underlying cell motility.

<table><tbody><tr><td>10 cm cell culture plates</td><td>Greiner</td><td>P7612-360EA</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Sigma-Aldrich</td><td>A7906</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s modified Eagle medium (DMEM)</td><td>Biological Industries, Israel</td><td>01-055-1A</td><td>Medium contains high glucose (4.5 g/L D-glucose)</td></tr><tr><td>Dulbecco&amp;rsquo;s phosphate buffered saline (1xDPBS)</td><td>Biological Industries, Israel</td><td>02-023-1A</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Biological Industries, Israel</td><td>04-001-1A</td><td></td></tr><tr><td>Fibronectin from human plasma, liquid, 0.1%, suitable for cell culture</td><td>Sigma-Aldrich</td><td>F0895</td><td></td></tr><tr><td>Glass bottom dishes</td><td>Cellvis</td><td>D35-20-1.5-N</td><td>35mm glass bottom dish, dish size 35 mm, well size 20mm, #1.5 cover glass (0.16-0.19 mm).</td></tr><tr><td>ImageJ software</td><td>NIH</td><td></td><td>Feely available at: https://imagej.nih.gov/ij/download.html</td></tr><tr><td>LAS-AF Leica Application Suite 3.2</td><td></td><td></td><td>Microscope acquisition software equipped with an ORCA-Flash 4.0 V2 digital CMOS</td></tr><tr><td>Leica AF6000</td><td>Leica</td><td></td><td>Inverted bright field microscope (40x, NA 1.3 ) equipped with phase-contrast optics, an incubator, and CO2 unit with LAS AF acquisition software equipped with an ORCA-Flash 4.0 V2 digital CMOS camera .</td></tr><tr><td>L-glutamine solution</td><td>Biological Industries, Israel</td><td>03-020-1B</td><td></td></tr><tr><td>ORCA-Flash 4.0 V2 digital CMOS camera</td><td>Hamamatsu Photonics</td><td></td><td></td></tr><tr><td>Penicillin-streptomycin solution</td><td>Biological Industries, Israel</td><td>03-031-1B</td><td></td></tr><tr><td>Trypsin-EDTA solution B (0.25%), EDTA (0.05%)</td><td>Biological Industries, Israel</td><td>03-052-1A</td><td></td></tr></tbody></table>

measuring cell-edge protrusion dynamics during spreading using live-cell microscopy

The development and homeostasis of multicellular organisms rely on coordinated regulation of cell migration. Cell migration is an essential event in the construction and regeneration of tissues, and is critical in embryonic development, immunological responses, and wound healing. Dysregulation of cell motility contributes to pathological disorders, such as chronic inflammation and cancer metastasis. Cell migration, tissue invasion, axon, and dendrite outgrowth all initiate with actin polymerization-mediated cell-edge protrusions. Here, we describe a simple, efficient, time-saving method for the imaging and quantitative analysis of cell-edge protrusion dynamics during spreading. This method measures discrete features of cell-edge membrane dynamics, such as protrusions, retractions, and ruffles, and can be used to assess how manipulations of key actin regulators impact cell-edge protrusions in diverse contexts.

In the experiment described in Figure 2, immortalized MEFs were plated on glass-bottom dishes pre-coated with fibronectin to activate integrin-mediated signaling, blocked by denatured BSA, to block free potential sites for cell adhesion which is not dependent on integrin activation. To reach the logarithmic growth phase at 70%-80% confluence of cells on the day of the experiment, 0.7 x 106 MEFs were plated in a 10 cm diameter tissue culture plate 16 h before the experiment. On the...

Watch this Scientific Journal Video about Measuring Cell-Edge Protrusion Dynamics during Spreading using Live-Cell Microscopy at JoVE.com

Measuring Cell-Edge Protrusion Dynamics during Spreading using Live-Cell Microscopy

This protocol aims to measure the dynamic parameters (protrusions, retractions, ruffles) of protrusions at the edge of spreading cells.

The development and homeostasis of multicellular organisms rely on coordinated regulation of cell migration. Cell migration is an essential event in the ...

measuring-cell-edge-protrusion-dynamics-during-spreading-using-live

Research

JoVE Journal

Biology

2.3K Views.  Bar-Ilan University. This protocol aims to measure the dynamic parameters (protrusions, retractions, ruffles) of protrusions at the edge of spreading cells.

Video: Measuring Cell-Edge Protrusion Dynamics during Spreading using Live-Cell Microscopy

Using the Dot Assay to Analyze Migration of Cell Sheets

Although complex organisms appear static, their tissues are under a continuous turnover. As cells age, die, and are replaced by new cells, cells move within tissues in a tightly orchestrated manner. During tumor development, this equilibrium is disturbed, and tumor cells leave the epithelium of origin to invade the local microenvironment, to travel to distant sites, and to ultimately form metastatic tumors at distant sites. The dot assay is a simple, two-dimensional unconstrained migration assay, to assess the net movement of cell sheets into a cell-free area, and to analyze parameters of cell migration using time-lapse imaging. Here, the dot assay is demonstrated using a human invasive, lung colony forming breast cancer cell line, MCF10CA1a, to analyze the cells' migratory response to epidermal growth factor (EGF), which is known to increase malignant potential of breast cancer cells and to alter the migratory phenotype of cells.

Cell migration is essential for development, tissue maintenance and repair, and tumorigenesis, and is regulated by growth factors, chemokines, and cytokines. This protocol describes the dot assay, a two-dimensional, unconstrained migration assay to assess the migratory phenotype of attached, cohesive cell sheets in response to microenvironmental cues.

Although complex organisms appear static, their tissues are under a continuous turnover. As cells age, die, and are replaced by new cells, cells move ...

Cancer Research

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Bioengineering

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Developmental Biology

Examining the Dynamics of Cellular Adhesion and Spreading of Epithelial Cells on Fibronectin During Oxidative Stress

The adhesion and spreading of cells onto the extracellular matrix (ECM) are essential cellular processes during organismal development and for the homeostasis of adult tissues. Interestingly, oxidative stress can alter these processes, thus contributing to the pathophysiology of diseases such as metastatic cancer. Therefore, understanding the mechanism(s) of how cells attach and spread on the ECM during perturbations in redox status can provide insight into normal and disease states. Described below is a step-wise protocol that utilizes an immunofluorescence-based assay to specifically quantify cell adhesion and spreading of immortalized fibroblast cells on fibronectin (FN) in vitro. Briefly, anchorage-dependent cells are held in suspension and exposed to the ATM kinase inhibitor Ku55933 to induce oxidative stress. Cells are then plated on FN-coated surface and allowed to attach for predetermined periods of time. Cells that remain attached are fixed and labeled with fluorescence-based antibody markers of adhesion (e.g., paxillin) and spreading (e.g., F-actin). Data acquisition and analysis are performed using commonly available laboratory equipment, including an epifluorescence microscope and freely available Fiji software. This procedure is highly versatile and can be modified for a variety of cell lines, ECM proteins, or inhibitors in order to examine a broad range of biological questions.

This method is useful for quantifying the early dynamics of cellular adhesion and spreading of anchorage-dependent cells onto the fibronectin. Furthermore, this assay can be used to investigate the effects of altered redox homeostasis on cell spreading and/or cell adhesion-related intracellular signaling pathways.

The adhesion and spreading of cells onto the extracellular matrix (ECM) are essential cellular processes during organismal development and for the ...

Biochemistry

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial cells regulate permeability of the endothelium, and many studies have indicated the important contribution of the actin cytoskeleton to determining junctional integrity1-5. A cortical actin belt is thought to be important for the maintenance of stable junctions1, 2, 4, 5. In contrast, actin stress fibers are thought to generate centripetal tension within endothelial cells that weakens junctions2-5. Much of this theory has been based on studies in which endothelial cells are treated with inflammatory mediators known to increase endothelial permeability, and then fixing the cells and labeling F-actin for microscopic observation. However, these studies provide a very limited understanding of the role of the actin cytoskeleton because images of fixed cells provide only snapshots in time with no information about the dynamics of actin structures5. 
Live-cell imaging allows incorporation of the dynamic nature of the actin cytoskeleton into the studies of the mechanisms determining endothelial barrier integrity. A major advantage of this method is that the impact of various inflammatory stimuli on actin structures in endothelial cells can be assessed in the same set of living cells before and after treatment, removing potential bias that may occur when observing fixed specimens. Human umbilical vein endothelial cells (HUVEC) are transfected with a GFP-&beta;-actin plasmid and grown to confluence on glass coverslips. Time-lapse images of GFP-actin in confluent HUVEC are captured before and after the addition of inflammatory mediators that elicit time-dependent changes in endothelial barrier integrity. These studies enable visual observation of the fluid sequence of changes in the actin cytoskeleton that contribute to endothelial barrier disruption and restoration. 
Our results consistently show local, actin-rich lamellipodia formation and turnover in endothelial cells. The formation and movement of actin stress fibers can also be observed. An analysis of the frequency of formation and turnover of the local lamellipodia, before and after treatment with inflammatory stimuli can be documented by kymograph analyses. These studies provide important information on the dynamic nature of the actin cytoskeleton in endothelial cells that can used to discover previously unidentified molecular mechanisms important for the maintenance of endothelial barrier integrity.

Microscopic imaging of live endothelial cells expressing GFP-actin allows characterization of dynamic changes in cytoskeletal structures. Unlike techniques that use fixed specimens, this method provides a detailed assessment of temporal changes in the actin cytoskeleton in the same cells before, during, and after various physical, pharmacological, or inflammatory stimuli.

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...

Lens-free Video Microscopy for the Dynamic and Quantitative Analysis of Adherent Cell Culture

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator and that it is possible to monitor and quantify single cells along several cell cycles. We describe the full protocol used to monitor and quantify a HeLa cell culture for 2.7 days. First, cell culture acquisition is performed with a lens-free video microscope, and then the data is analyzed following a four-step process: multi-wavelength holographic reconstruction, cell-tracking, cell segmentation and cell division detection algorithms. As a result, we show that it is possible to gather a dataset featuring more than 10,000 cell cycle tracks and more than 2 x 106 cell morphological measurements.

Lens-free video microscopy enables us to monitor cell cultures directly inside the incubator. Here we describe the full protocol used to acquire and analyze a 2.7 day long acquisition of cultured HeLa cells, leading to a dataset of 2.2 x 106 measurements of individual cell morphology and 10584 cell cycle tracks.

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction forces have been well-characterized, but there is a lack of techniques allowing the measurement of the protrusion forces exerted by cells orthogonally to their substrate. We designed an experimental setup to measure the protrusion forces exerted by adherent cells on their substrate. Cells plated on a compliant Formvar sheet deform this substrate and the resulting topography is mapped by atomic force microscopy (AFM) at the nanometer scale. Force values are then extracted from an analysis of the deformation profile based on the geometry of the protrusive cellular structures. Hence, the forces exerted by the individual protruding units of a living cell can be measured over time. This technique will enable the study of force generation and its regulation in the many cellular processes involving protrusion. Here, we describe its application to measure the protrusive forces generated by podosomes formed by human macrophages.

Here, we detail the experimental techniques used to evaluate the protrusion forces that podosomes apply on a compliant film, from the preparation of the film to the automated analysis of topographical images.

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events

Durotaxis is the process by which cells sense and respond to gradients of tension. In order to study this process in vitro, the stiffness of the substrate underlying a cell must be manipulated. While hydrogels with graded stiffness and long-term migration assays have proven useful in durotaxis studies, immediate, acute responses to local changes in substrate tension allow focused study of individual cell movements and subcellular signaling events. To repeatably test the ability of cells to sense and respond to the underlying substrate stiffness, a modified method for application of acute gradients of increased tension to individual cells cultured on deformable hydrogels is used which allows for real time manipulation of the strength and direction of stiffness gradients imparted upon cells in question. Additionally, by fine tuning the details and parameters of the assay, such as the shape and dimensions of the micropipette or the relative position, placement, and direction of the applied gradient, the assay can be optimized for the study of any mechanically sensitive cell type and system. These parameters can be altered to reliably change the applied stimulus and expand the functionality and versatility of the assay. This method allows examination of both long term durotactic movement as well as more immediate changes in cellular signaling and morphological dynamics in response to changing stiffness.

Mechanical forces are important for controlling cell migration. This protocol demonstrates the use of elastic hydrogels that can be deformed using a glass micropipette and a micromanipulator to stimulate cells with a local stiffness gradient to elicit changes in cell structure and migration.

Durotaxis is the process by which cells sense and respond to gradients of tension. In order to study this process in vitro, the stiffness of the substrate ...

Measuring Cell-Edge Protrusion Dynamics during Spreading using Live-Cell Microscopy

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Chapters in this video

Using the Dot Assay to Analyze Migration of Cell Sheets

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Examining the Dynamics of Cellular Adhesion and Spreading of Epithelial Cells on Fibronectin During Oxidative Stress

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Lens-free Video Microscopy for the Dynamic and Quantitative Analysis of Adherent Cell Culture

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events