Name: measuring cell-edge protrusion dynamics during spreading using live-cell microscopy
Uploaded: 2021-11-01T13:00:00
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 C5...

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#8217;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&#8217;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 ...

The cell-edge protrusion assay has been shown to directly correlate with cell migration. Therefore it can be used as a preliminary method for identifying critical proteins and signaling mechanisms involved in cell motility. The method is fast, simple, cost effective and does not require fluorescent labeling or using an expensive fluorescent microscope.Demonstrating the procedure will be Michal Gendler, a student from my laboratory. Add two milliliters of one normal hydrochloric acid solution at the center of a glass bottom dish and incubate for 20 minutes at room temperature. Wash the dish three times with two milliliters of PBS.Dilute fibronectin at 10 micrograms per milliliter concentration in PBS and add 200 microliters of the diluted solution to the glass center dish. Incubate for one hour at 37 degrees Celsius. Prepare 1%BSA solution and PBS and pass it through a 0.2 micrometer filter.Denature the solution by incubating at 70 degrees Celsius for 30 minutes in a pre-warmed water bath. Wash the coated glass bottom dish with two milliliters of PBS three times. Add two milliliters of denatured BSA solution and incubate the dish at 37 degree Celsius for one hour.Wash the glass dish with two milliliters of PBS three times. Before 16 to 18 hours of experiment at 0.7 million cells per 10 centimeter diameter tissue culture plate to achieve 70 to 80%confluency. The next day, add two milliliters of trypsin solution to the tissue culture plate and incubate for two to three minutes until the cells get detached.Add five milliliters of the medium to inactivate the trypsin. Using a hemacytometer, count the cells and plate 20, 000 cells in two milliliters of the complete medium in a bottom dish. Then incubate the dish with plated cells in an incubator for 15 minutes.Turn on the heating unit and set it at 37 degrees Celsius one hour before imaging. Also, turn on the carbon dioxide unit and set it at 5%10 minutes before imaging. Switch on the microscope and camera.Turn on the computer and open the microscope acquisition software. Set the magnification on a 40X dry lens phase contrast. Set the total movie duration to 10 minutes with a time interval of five seconds.After 15 minutes of incubation, place the glass bottom dish with adhered cells into an adapter and fix it. Insert the adapter with the dish into the slots in the microscope stage. Take off the dish cover, place the carbon dioxide lid and open the carbon dioxide valve.Locate an appropriate cell for imaging. Start movie acquisition after focusing on the cell. To perform the image analysis, select the straight tool in the image analysis software and make eight lines of 20 arbitrary units perpendicular to the protrusions in a radial arrangement at every 45 degrees, including lamella and cell-edge.In the main toolbar, go to image, select Stacks and then click on Reslice to generate a kymograph picture describing the movement of single points within the cell membrane. Using the kymograph images, extract and manually count 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 minutes.Determine the protrusion persistence, distance and velocity by kymography analysis. To determine the protrusion distance, draw perpendicular line from the base of the protrusion to the highest peak of the protrusion. Press M to measure the length of the line in pixels and ensure that the pixel to micrometer ratio is known to convert the length in micrometers.Quantification of protrusions, reactions, and ruffles was performed manually per 10 minutes and the average frequency obtained for protrusions and retractions was 5.1 and 2.1 for ruffles. The representative kymograph had a protrusion distance of approximately 4.8 micrometers and a protrusion time of approximately 0.6 minutes. An example of a cell that should be excluded from the analysis is represented.The kymography analysis revealed that the cell was not present in its spreading phase and did not show any clear membrane protrusions. The most important thing is to choose correct cells for imaging. A proper cell should be in its spreading phase and should not touch other cells to avoid cell-cell communication and signaling.The method can be used as a preliminary tool for testing cytoskeletal dynamics involving cell motility before deciding to perform more results demanding cell migration assays.

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Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to investigate cells behavior at sub-cellular level with high spatial and temporal resolution was developed. Thus, an atomic force microscope (AFM) was integrated with total internal reflection fluorescence (TIRF) microscopy and fast-spinning disk (FSD) confocal microscopy. The integrated system is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells, allowing direct optical imaging of cell responses to mechanical stimulation in real-time. Significant rearrangement of the actin filaments and focal adhesions was shown due to local mechanical stimulation at the apical cell surface that induced changes into the cellular structure throughout the cell body. These innovative techniques will provide new information for understanding live cell restructuring and dynamics in response to mechanical force. A detailed protocol and a representative data set that show live cell response to mechanical stimulation are presented.

This paper aims to instruct the reader in the operation of an integrated atomic force-optical imaging microscope for mechanical stimulation of live cells in culture. A step-by-step protocol is presented. A representative data set that shows live cell response to mechanical stimulation is presented.

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single procedure. Due to advances in microscope technology it is relatively easy to capture live cell images for the purpose of investigating cellular events in real time with minimal concern regarding phototoxicity to the cells. This protocol describes how to take live cell timelapse images of primary rat neonatal cardiomyocytes using a confocal spinning disk microscope following lentiviral and adenoviral transduction to modulate properties of the cell. The application of two different types of viruses makes it easier to achieve an appropriate transduction rate and expression levels for two different genes. Well focused live cell images can be obtained using the microscope&#8217;s autofocus system, which maintains stable focus for long time periods. Applying this method, the functions of exogenously engineered proteins expressed in cultured primary cells can be analyzed. Additionally, this system can be used to examine the functions of genes through the use of siRNAs as well as of chemical modulators.

This protocol describes a method of live cell imaging using primary rat neonatal cardiomyocytes following lentiviral and adenoviral transduction using confocal spinning disk microscopy. This enables detailed observations of cellular processes in living cardiomyocytes.

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Live Cell Imaging and 3D Analysis of Angiotensin Receptor Type 1a Trafficking in Transfected Human Embryonic Kidney Cells Using Confocal Microscopy

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in transfected human embryonic kidney-293 (HEK) cells following stimulation with angiotensin II (Ang II). HEK cells are transiently transfected with plasmid DNA containing AT1aR tagged with enhanced green fluorescent protein (EGFP). Lysosomes are identified with a red fluorescent dye. Live-cell images are captured on a laser scanning confocal microscope after Ang II stimulation and analyzed by software in three dimensions (3D, voxels) over time. Live-cell imaging enables investigations into receptor trafficking and avoids confounds associated with fixation, and in particular, the loss or artefactual displacement of EGFP-tagged membrane receptors. Thus, as individual cells are tracked through time, the subcellular localization of receptors can be imaged and measured. Images must be acquired sufficiently rapidly to capture rapid vesicle movement. Yet, at faster imaging speeds, the number of photons collected is reduced. Compromises must also be made in the selection of imaging parameters like voxel size in order to gain imaging speed. Significant applications of live-cell imaging are to study protein trafficking, migration, proliferation, cell cycle, apoptosis, autophagy and protein-protein interaction and dynamics, to name but a few.

Here we present a protocol to image cells expressing green fluorescent protein-tagged angiotensin type 1a receptors during endocytosis initiated by angiotensin II treatment. This technique includes labeling lysosomes with a second fluorescent marker, and then utilizing software to analyze the co-localization of receptor and lysosome in three dimensions over time.

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

Plasma sterilization is a promising alternative to conventional sterilization methods for industrial, clinical, and spaceflight purposes. Low pressure plasma (LPP) discharges contain a broad spectrum of active species, which lead to rapid microbial inactivation. To study the efficiency and mechanisms of sterilization by LPP, we use spores of the test organism Bacillus subtilis because of their extraordinary resistance against conventional sterilization procedures. We describe the production of B. subtilis spore monolayers, the sterilization process by low pressure plasma in a double inductively coupled plasma reactor, the characterization of spore morphology using scanning electron microscopy (SEM), and the analysis of germination and outgrowth of spores by live cell microscopy. A major target of plasma species is genomic material (DNA) and repair of plasma-induced DNA lesions upon spore revival is crucial for survival of the organism. Here, we study the germination capacity of spores and the role of DNA repair during spore germination and outgrowth after treatment with LPP by tracking fluorescently-labelled DNA repair proteins (RecA) with time-resolved confocal fluorescence microscopy. Treated and untreated spore monolayers are activated for germination and visualized with an inverted confocal live cell microscope over time to follow the reaction of individual spores. Our observations reveal that the fraction of germinating and outgrowing spores is dependent on the duration of LPP-treatment reaching a minimum after 120 s. RecA-YFP (yellow fluorescence protein) fluorescence was detected only in few spores and developed in all outgrowing cells with a slight elevation in LPP-treated spores. Moreover, some of the vegetative bacteria derived from LPP-treated spores showed an increase in cytoplasm and tended to lyse. The described methods for analysis of individual spores could be exemplary for the study of other aspects of spore germination and outgrowth.

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

This protocol illustrates the important consecutive steps required to assess the relevance of monitoring vitality parameter and DNA repair processes in reviving Bacillus subtilis spores after treatment with low pressure plasma by tracking fluorescence-labelled DNA repair proteins via time-resolved confocal microscopy and scanning electron microscopy.

Plasma sterilization is a promising alternative to conventional sterilization methods for industrial, clinical, and spaceflight purposes. Low pressure ...

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 ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear positioning occurs during skeletal muscle differentiation. Muscle fibers (myofibers) are multinucleated cells formed by the fusion of muscle precursor cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation. Correct nuclear positioning within myofibers is required for the proper muscle regeneration and function. The common procedure to assess myoblast differentiation and myofiber formation relies on fixed cells analyzed by immunofluorescence, which impedes the study of nuclear movement and cell behavior over time. Here, we describe a method for the analysis of myoblast differentiation and myofiber formation by live cell imaging. We provide a software for automated nuclear tracking to obtain a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior (i.e., the trajectory) during differentiation and fusion.

Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear ...