This work was funded by a National Health and Medical Research Council (NHMRC) Project Grant 568721 (SRT) and, in part, by a UNSW Faculty of Medicine Faculty Research Grant (MDR).

1. General Endothelial Cell Culture
<ol>
	<li>Culture bovine aortic endothelial cells (passages 4&#8211;9) on gelatin-coated tissue culture flasks (coat tissue culture surface with 0.1% w/v gelatin in PBS at RT for 15 min) in EGM-2 media (with the EGM-2 bullet kit containing 5% fetal bovine serum, growth factors and all supplements provided by the manufacturer, except for hydrocortisone).</li>
	<li>When cells are near-confluent (ca. 3 days post seeding after a 1:4 split), harvest cells by treatment with 0.05% w/v trypsin / 0.02% w/v EDTA in PBS at 37 &#176;C. After the majority of cells have detached, add complete EGM-2 media to quench the trypsin and then centrifuge (100 x g, 5 min).</li>
	<li>Prepare cells for studies on the de novo adhesion of endothelial cells to fibronectin (Experiment 1: Section 2) and the subsequent de-adhesion of endothelial cells with established adhesion on this substrate (Experiment 2: Section 3).
	<ol>
		<li>Wash harvested cells once with serum-free Medium 199 containing 1% w/v bovine serum albumin (BSA) and re-centrifuge (100 x g, 5 min).</li>
		<li>Re-suspend cells in serum-free Medium 199 containing 1% w/v BSA at 2.5 x 105 cells/ml (cell-substrate impedance measurements) or 5 x 105 cells/ml (live cell imaging analyses) and maintain at 37 &#176;C prior to use. 
		NOTE: The adhesion responses of cells are highly sensitive to temperature differences (e.g., due to convection effects) so all equipment and solutions used to handle and treat cells during the following protocols should be kept at a constant temperature of 37 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Experiment 1: Quantifying De Novo Endothelial Cell Adhesion on Native and MPO-oxidized Fibronectin (Cell-substrate Impedance)
NOTE: Experiment 1 examines the degree to which MPO-mediated oxidation of fibronectin impairs its ability to support de novo adhesion of suspended endothelial cells.
<ol>
	<li>Coat fibronectin onto 96-well gold cell-substrate impedance microelectrode arrays. Add 80 &#956;l/well of purified bovine fibronectin at 5 &#956;g/ml in PBS, incubate for 2 hr at 37 &#176;C and remove the solution.</li>
	<li>Incubate fibronectin-coated surfaces with MPO to allow the binding of MPO to the surface bound fibronectin. Add 80 &#956;l/well of purified human neutrophil MPO at 20 nM in Hank&#8217;s balanced salt solution (HBSS) and incubate for 0.5 hr at 37 &#176;C.</li>
	<li>Wash surfaces twice with HBSS to remove any unbound MPO.</li>
	<li>Add H2O2 (0-10 &#956;M final concentration) to wells of the microelectrode array plate containing 80 &#956;l/well HBSS to initiate MPO-catalyzed, HOCl-dependent fibronectin oxidation and incubate for another 0.5 hr at 37 &#176;C.</li>
	<li>To examine the effect of relevant inhibitors or modulators of MPO-catalyzed reactions (e.g., alternative MPO enzyme substrates, enzyme inhibitors or antioxidants; see9 for details), add these to the HBSS immediately prior to H2O2 addition.</li>
	<li>After 0.5 hr, treat surfaces with methionine to quench residual surface-bound oxidizing species; i.e., reactive protein-bound chloramines, which may exert confounding cellular activities. Add 10 mM methionine in 80 &#956;l HBSS per well and incubate for 10 min at 37 &#176;C.</li>
	<li>Block surfaces with BSA. Add 80 &#956;l/well of BSA at 0.2% w/v in PBS, incubate for 2 hr at 37 &#176;C and remove the solution. 
	NOTE: Residual uncoated surface regions will support cell adhesion and blocking these with a non-adhesive protein (i.e., BSA) ensures that cellular adhesion responses strictly depend on the purified cell-adhesive matrix employed, in this case fibronectin.</li>
	<li>Wash surfaces twice with HBSS. 
	NOTE: None of the preceding surface treatments appreciably affects cell-substrate readings.</li>
	<li>Seed suspended endothelial cells (add 200 &#956;l/well at ca. 2.5 x 105 cells/ml, prepared in serum-free Medium 199 containing 1% BSA; see 1.3) onto the native or MPO-oxidized fibronectin coated surfaces.</li>
	<li>Immediately after seeding cells (i.e., prior to any cell attachment and spreading), mount the microelectrode array plate onto the incubator port (housed in a 37 &#176;C incubator in the presence of 5% CO2).
	<ol>
		<li>Using the instrument software immediately take a &#8216;blank&#8217; reading to normalize subsequent cell-substrate impedance (&#8216;cell index&#8217;) values to the initial background values obtained in the absence of cell-adhesion.</li>
		<li>Initiate acquisition of continuous cell index data (minimum of one cell index reading/min).</li>
	</ol>
	</li>
	<li>Incubate cells at 37 &#176;C and 5% CO2 for 2 hr, a time period during which maximal cell attachment and spreading is achieved; this is reflected by a plateauing of the cell index values (see Figure 3A). 
	NOTE: The preceding experiment (Experiment 1) examines how initial MPO-mediated oxidation of fibronectin limits the ability of endothelial cells to establish cell adhesion on this substrate. The following experiment (Experiment 2) examines how MPO-mediated fibronectin oxidation promotes decreases in cell-matrix adhesion (i.e., de-adhesion) in endothelial cells with established adhesion onto this substrate. The treatments in these two experiments are essentially identical, except for the timing of MPO-mediated fibronectin oxidation (i.e., before cell adhesion &#8211; Experiment 1; after cell adhesion &#8211; Experiment 2).</li>
</ol>
3. Experiment 2: Quantifying Endothelial Cell De-adhesion from Fibronectin in Response to MPO-mediated Fibronectin Oxidation (Cell-substrate Impedance and Live Cell Imaging)
<ol>
	<li>Coat fibronectin onto 96-well gold microelectrode arrays for cell-substrate impedance measurements (add 80 &#956;l/well of fibronectin at 5 &#956;g/ml in PBS and incubate for 2 hr at 37 &#176;C) or 35 mm glass-bottomed cell culture dishes for live cell imaging analyses (add 2 ml/dish of fibronectin at 5 &#956;g/ml in PBS and incubate for 2 hr at 37 &#176;C).</li>
	<li>Block surfaces with BSA. Add BSA at 0.2% w/v in PBS at the volumes indicated in 3.1 and incubate for 2 hr at 37 &#176;C.</li>
	<li>Incubate surfaces with MPO to allow the binding of MPO to fibronectin. Add 20 nM purified human MPO in HBSS at the volumes indicated in 3.1 and incubate for 0.5 hr at 37 &#176;C.</li>
	<li>Wash surfaces twice with HBSS to remove any unbound MPO.</li>
	<li>Seed suspended endothelial cells (prepared in serum-free Medium 199 containing 1% w/v BSA; see 1.3) onto the native or MPO-bearing fibronectin coated surfaces.
	<ol>
		<li>Add 200 &#956;l/well at 2.5 x 105 cells/ml to 96 well cell-substrate impedance microelectrode arrays.</li>
		<li>Add 2 ml/dish at 5 x 105 cells/ml to 35 mm glass bottomed cell culture dishes.</li>
	</ol>
	</li>
	<li>Immediately after seeding cells (i.e., prior to any cell attachment and spreading):
	<ol>
		<li>Mount 96 well microelectrode array plates onto the cell-substrate impedance incubator port (housed in a 37 &#176;C incubator in the presence of 5% CO2) and take &#8216;blank&#8217; readings to initiate continuous acquisition of cell index data (cf. Section 2.10).</li>
		<li>Transfer 35 mm glass bottomed cell culture dishes to a 37 &#176;C incubator in the presence of 5% CO2.</li>
	</ol>
	</li>
	<li>Incubate cells at 37 &#176;C for 2 hr to allow maximal cell attachment and spreading (cf. Section 2.11).</li>
	<li>Briefly remove the 96 well microelectrode array plate (pause cell index readings at this point) or 35 mm glass bottomed cell culture dish from the 37 &#176;C incubator, remove cell supernatant and add warmed (37 &#176;C) HBSS (volumes as per step 3.1). 
	NOTE: HBSS is employed when studying MPO-catalyzed oxidative reactions instead of complete culture media as the latter contains oxidizable species that interfere with the oxidation reactions.</li>
	<li>To examine the effect of inhibitors/modulators of MPO-catalyzed reactions (alternative enzyme substrates, enzyme inhibitors or antioxidants) or cell signaling inhibitors (e.g., 40 &#956;M blebbistatin to inhibit actomyosin contractility), add these now.</li>
	<li>Immediately place the microelectrode array plate back into the 37 &#176;C incubator port (re-commence cell index readings at this point) or the 35 mm glass bottomed cell culture dish back into the 37 &#176;C incubator, and allow the cells to equilibrate in the HBSS for 0.5 hr. 
	NOTE: After 0.5 hr equilibration in HBSS, cell index values stabilize at slightly lower values; when pre-incubating cells with enzyme substrates, enzyme and cell signaling inhibitors or antioxidants, first ensure that these do not significantly affect the values obtained after equilibration.</li>
	<li>Initiate MPO-catalyzed, HOCl-dependent fibronectin oxidation and de-adhesion.
	<ol>
		<li>For cell impedance studies using 96 well microelectrode array plates:
		<ol>
			<li>Remove the microelectrode array plate from the incubator and pause cell index measurements.</li>
			<li>Add H2O2 (final concentration of 0-10 &#956;M) to the HBSS and gently mix by repeated pipetting.</li>
			<li>Immediately, re-mount the microelectrode array back onto the 37 &#176;C incubator port and re-commence the acquisition of cell index readings.</li>
		</ol>
		</li>
		<li>For live cell imaging studies using 35 mm glass bottomed cell culture dish:
		<ol>
			<li>Remove the culture dish from the incubator and mount onto a 37 &#176;C heated stage of an inverted confocal microscope equipped with a 63X water objective lens with suitable DIC optics for recording live cell movies.</li>
			<li>Focus on cells and optimize DIC optics (K&#246;hler illumination, bias retardation and camera offset/gain; for details, see13).</li>
			<li>Initiate DIC movie and record baseline readings of untreated cells for 1 min.</li>
			<li>Add H2O2 (final concentration of 0-10 &#956;M) and gently mix by repeated pipetting. Re-focus the microscope (if necessary) and continue recording DIC movies of the treated cells for the required time period.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Data Analysis and Presentation
<ol>
	<li>Cell-substrate impedance data
	<ol>
		<li>Export raw data (cell index versus time) into a spreadsheet.</li>
		<li>For cell de-adhesion studies (Experiment 2: Section 3), normalize data by setting values recorded immediately prior to the initiation of MPO-mediated fibronectin oxidation at a value of 1 (i.e., immediately before the addition of H2O2 in 3.11.1.1). 
		NOTE: This ensures that relative changes in cell index values elicited by MPO-mediated fibronectin oxidation are not masked by small initial differences in absolute cell index values between wells.
		<ol>
			<li>Present data as plots of normalized cell index (y-axis) versus time (x-axis).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Live cell imaging data (cell de-adhesion studies in Experiment 2: Section 3)
	<ol>
		<li>Open DIC live cell imaging movies recorded immediately prior to and after the initiation of MPO-mediated fibronectin oxidation in a standard image analysis program (e.g., ImageJ software).</li>
		<li>In at least two separate DIC movies randomly select multiple cells and measure their projected area in sequential frames (e.g., at 1 min intervals) by manually tracing their membrane edge and quantifying the number of enclosed pixels.</li>
		<li>Export raw data (projected cell area versus time) to an Excel spreadsheet and normalize cell area data by setting values recorded immediately prior to the initiation of MPO-mediated fibronectin oxidation at a value of 1 (i.e., immediately before the addition of H2O2 in 3.11.2.3).</li>
		<li>Present data as a plot of normalized cell area (y-axis) versus time (x-axis).</li>
	</ol>
	</li>
</ol>

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E., Schiller, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+Interference+Contrast+(DIC)+Imaging+of+Living+Cells.">Differential Interference Contrast (DIC) Imaging of Living Cells.</a> CSH Protoc. 2007, (2007).</li><li>Kennett, E. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peroxynitrite+modifies+the+structure+and+function+of+the+extracellular+matrix+proteoglycan+perlecan+by+reaction+with+both+the+protein+core+and+the+heparan+sulfate+chains.">Peroxynitrite modifies the structure and function of the extracellular matrix proteoglycan perlecan by reaction with both the protein core and the heparan sulfate chains.</a> Free Radic Biol Med. 49, 282-293 (2010).</li><li>Sen, S., Ng, W. 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The authors have no disclosures to make.

Cell-matrix adhesion and de-adhesion processes can be quantified accurately in real time with high temporal resolution using cell-substrate impedance and live cell imaging analyses. These real-time approaches provide a major advantage over end-point analyses of cell adhesion, which provide poor temporal resolution. By accurately quantifying rapid de-adhesion responses with high temporal resolution, these analyses can provide critical insights into how morphological responses to matrix modifications are regulated and how they impact on adhesion-dependent cell signaling processes, measured in parallel using relevant biochemical assays (e.g., Western blotting).
In recent studies, these approaches were used to reveal a novel role for MPO-mediated oxidation of the subendothelial matrix in triggering the de-adhesion of endothelial cells and showed: (i) that the rate of de-adhesion was critically dependent on pre-existing actomyosin contractility (cf. Figures 3B and 3C) and (ii) that the loss of cell-matrix adhesion drove changes in important adhesion-dependent cell signaling pathways including Src-dependent paxillin phosphorylation and myosin light chain II phosphorylation9 (see Figure 1). These data have important implications for understanding endothelial function and barrier integrity during inflammatory responses, where the subendothelial extracellular matrix is implicated as a key target for oxidants produced by subendothelial deposits of matrix-bound MPO or other sources of reactive oxidants 3,5-9,14.
The use of a microarray-based, cell-substrate impedance biosensor system provides a platform for robust, high-throughput cell adhesion measurements. Each well of the 96 well cell-substrate impedance microelectrode arrays has the potential to simultaneously analyze 32 different experimental conditions (i.e., 3 wells per condition). However, when comparing small changes in cell adhesion, at least 4 or more wells should be used per experimental condition. During cell seeding and subsequent cell treatments, care should be taken to keep all solutions at 37 &#176;C and minimize the time taken to treat cells outside the 37 &#176;C incubator environment (this avoids potential temperature differences across the microelectrode array that may affect adhesion responses). Notably, the electrical current at the cell-substrate interface is sensitive to the ionic strength of the cell supernatant10: consequently, cells should be allowed to equilibrate after the addition of new buffers/media to obtain a steady cell index response before monitoring the effect of the stimulus of interest (see 3.10 and Figure 3A). Decreases in cell index may not only reflect a decrease in the average area of cell-matrix contact but may also reflect a decrease in the number of attached cells. To discriminate between these possibilities, changes in the number of attached cells on microelectrode arrays can be quantified immediately after cell-substrate impedance measurements by Crystal violet staining this approach has been used to confirm that MPO-mediated fibronectin oxidation triggers a rapid decrease in the average area of cell-matrix contact, but does not promote cell detachment (for details, see9).
A limitation of the cell-substrate impedance technique is that it provides an average measure of adhesion changes over all cells present on the microelectrode surface and it does not give information about the precise nature of adhesion or morphological changes at the level of individual cells. To address this limitation, live cell DIC microscopy and image analysis provides a complementary measure of adhesion changes and reveals morphological changes of individual cells. Thus, decreases in cell index in response to MPO-mediated fibronectin oxidation measured by cell-substrate impedance (Figure 3A) correlate well with the decreases in the projected area of cells measured by live cell image analysis of DIC movies (Figure 3B). Importantly, DIC movies reveal that, in individual cells, MPO-induced de-adhesion involves the rapid retraction of the peripheral cell membrane from the substratum and adjacent cells and ultimately results in cells assuming a compact &#8216;rounded-up&#8217; morphology, but without resulting in cell detachment (Movie 1). As with cell-substrate impedance measurements, to ensure reproducible cellular responses, care should be taken to maintain solutions at 37 &#176;C prior to use on cells and a heated microscope stage (or equivalent temperature control device) must be employed during the live cell imaging recordings.
The protocols described here can be modified readily by substituting fibronectin with other purified matrix substrates (e.g., collagen or laminin) or studying a more complex matrix environment produced overtime by cultured adherent cells (for details, see9). The protocols can also be altered by substituting MPO-mediated matrix oxidation with other soluble stimuli that disrupt cell-matrix adhesion including other physiological matrix-modifying oxidants (e.g., peroxynitrite, nitrogen dioxide radical5-8), matrix-degrading proteases15 or antagonists of adhesive ligands present on the cell surface or in the matrix (e.g., anti-integrin antibodies or RGD peptides). Live cell imaging analyses can also be used to quantify cell-matrix de-adhesion in response to the electrochemical desorption of cell-matrix contacts16. Finally, the described protocols are also readily applicable to studying cellular adhesion dynamics in adherent cells other than endothelial cells (e.g., epithelial cells).

Stable adhesive contacts between cells and their surrounding extracellular matrix are required for maintenance of tissue homeostasis. For example, endothelial cell adhesion to the subendothelial matrix in blood vessels plays a critical role in maintaining the integrity of the endothelial layer and its homeostatic function as a regulatory, semi-permeable vascular barrier1. The actin cytoskeleton is mechanically coupled to adhesive matrix molecules at sites of cell-matrix adhesion and adhesive contacts at the cell-matrix interface play an important role in determining the position of the cell membrane by resisting centrally-directed actomyosin tensile forces. Extracellular stimuli that alter cell-matrix adhesion necessarily alter the balance of forces at the cell-matrix interface, an event that is rapidly &#8216;sensed&#8217; by mechano-sensitive signaling proteins, resulting in the transduction of &#8220;outside-in signaling&#8221;. This cross-talk between cells and their surrounding extracellular matrix plays a key role in controlling cell shape, motility, function, proliferation and survival2.
Diverse patho-physiological processes (embryonic development, inflammation, wound repair and cancer metastasis) are characterized by dynamic remodeling of adhesive matrix substrates by matrix-degrading oxidants and/or enzymes3,4. For example, adhesive subendothelial matrix proteins in blood vessels (e.g., fibronectin) are implicated as major targets for modification or degradation in human inflammatory diseases due to the localized production of reactive oxidants (e.g., hypochlorous acid, HOCl) by the leukocyte-derived enzyme myeloperoxidase (MPO), which accumulates within the subendothelium during inflammatory vascular disease (Figure 1)5-9. Changes in cell-matrix adhesion induced by MPO-derived oxidants and other matrix-modifying stimuli are likely to play important roles in altering vascular homeostasis during a variety of pathological processes; e.g., by altering endothelial cell signaling, morphology and viability, which in turn perturbs endothelial function and barrier integrity. However, the morphological and cell signaling responses of adherent cells to extracellular matrix modifications are only beginning to be understood.
To understand how matrix modifications drive changes in cell adhesion dynamics and adhesion-dependent cell signaling pathways, techniques are required that accurately quantify changes in cell-matrix adhesion in real time, with high temporal resolution. Here, we describe complementary cell-substrate impedance and live cell imaging techniques that fulfill these criteria and provide a platform to quantify cell adhesion and de-adhesion processes in a non-invasive manner.
We show how these cell-substrate impedance and live cell imaging approaches can be readily employed to (i) monitor the dynamics of cell attachment and spreading (i.e., de novo cell adhesion) onto native and modified matrix substrates and (ii) to measure the dynamics of cell-matrix detachment (i.e., de-adhesion) by adherent cells exposed to matrix-modifying stimuli. The xCELLigence cell-substrate impedance biosensor system provides a continuous measurement of the area of cell-matrix contact by quantifying electrical impedance at the surface of 96-well gold microelectrode arrays and expresses these electrical impedance measurements as &#8216;cell index&#8217;, a dimensionless value that is largely proportional to the area of cell-substrate contact10 (Figure 2), whilst also being sensitive to changes in the average distance between the (insulating) cell membrane and the electrode surface11. A further increase in cell index values is also achieved upon formation of tight cell-cell contacts that restrict paracellular current flows,11 conditions that do not prevail within the experiments described in this study. Measurement of the projected area of individual cells over time by image analysis of time-lapse differential interference contrast (DIC) movies provides a complementary measure of changes in the area of cell-substrate contact and provides additional information regarding the precise nature and dynamics of the morphological changes quantified by the cell-substrate impedance approach.
Specifically, we describe the application of these approaches to monitor how MPO-mediated oxidation of adhesive subendothelial matrix proteins (e.g., fibronectin) (i) reduces the de novo adhesion of suspended endothelial cells onto purified fibronectin and (ii) triggers cell-matrix de-adhesion in endothelial cells with established adhesion on fibronectin. By performing parallel cell signaling analyses over time using relevant biochemical assays (e.g., Western blotting), the temporal and causal relationships between adhesion/de-adhesion processes and associated changes in adhesion-dependent cell signaling events can be determined.
These approaches were recently used to demonstrate that extracellular matrix oxidation catalyzed by subendothelial deposits of MPO triggers a rapid loss in cell-matrix adhesion of endothelial cells that is driven by pre-existing actomyosin contractile forces9. Importantly, by enabling the temporal relationship between changes in both cell adhesion and adhesion-dependent cell signaling to be determined, these approaches identified that MPO-induced matrix modification and cellular de-adhesion triggers changes in important adhesion-dependent cell signaling pathways including Src kinase-dependent paxillin phosphorylation and myosin light chain II phosphorylation (Figure 1)9. This mode of redox-dependent signaling, involving the activation of intracellular signaling events by extracellular oxidative reactions that disrupt cell-matrix adhesion, represents a novel mode of cell signaling termed &#8220;outside-in redox signaling&#8221; (Figure 1)9.
In general, these complementary cell-substrate impedance biosensor and live cell imaging approaches should be valuable in revealing how different matrix-modifying stimuli or agents drive changes in cell adhesion dynamics, morphology and signaling within different adherent cell-types subject to a wide variety of experimental settings.
The following protocol describes how to quantify the impact of MPO-mediated matrix oxidation on de novo endothelial cell adhesion (Experiment 1) and endothelial cell de-adhesion (Experiment 2) processes. MPO binds avidly to fibronectin and other adhesive subendothelial extracellular matrix proteins and uses hydrogen peroxide (H2O2) to convert chloride ions (Cl&#8211;) to the highly reactive chlorinating oxidant hypochlorous acid (HOCl), which reacts locally with these matrix proteins and disrupts their cell adhesive properties (Figure 1)8,9,12.

<table><tbody><tr><td>96 well gold cell-substrate impedance microelectrode array</td><td>ACEA Biosciences / Roche</td><td>E-Plate 96</td><td>single-use plate used for performing cell-based assays on the xCELLigence system</td></tr><tr><td>blebbistatin</td><td>Sigma</td><td>B0560</td><td>selective inhibitor of non-muscle myosin-II</td></tr><tr><td>Bovine Aortic Endothelial Cells, cryopreserved&amp;nbsp;</td><td>Lonza</td><td>BW-6001</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma</td><td>05470</td><td></td></tr><tr><td>EGM-2 BulletKit</td><td>Lonza</td><td>CC-3162</td><td>endothelial cell growth media kit</td></tr><tr><td>Fibronectin, lyophilized powder</td><td>Sigma</td><td>F-4759</td><td>from bovine plasma</td></tr><tr><td>Fluorodish 35 mm glass-bottomed cell culture dish</td><td>World Precision Instruments</td><td>FD35-100</td><td>Cell cuture dish with optical quality glass bottom for imaging</td></tr><tr><td>Gelatin from bovine skin</td><td>Sigma</td><td>G9391</td><td>cell culture substratum</td></tr><tr><td>Hank&#039;s Balanced Salt Solution</td><td>Life Technologies</td><td>14025076</td><td></td></tr><tr><td>Hydrogen peroxide</td><td>Merck</td><td>107298</td><td></td></tr><tr><td>Medium-199</td><td>Life Technologies</td><td>11150-059</td><td>serum-free cell media</td></tr><tr><td>Methionine</td><td>Sigma</td><td>M9500</td><td>quenches chlorinating oxidants generated by myeloperoxidase</td></tr><tr><td>Myeloperoxidase, Human Polymorphonuclear Leukocytes</td><td>Millipore</td><td>475911</td><td></td></tr><tr><td>RTCA MP Instrument / xCELLigence cell substrate impedance system</td><td>ACEA Biosciences / Roche</td><td></td><td>Consists of RTCA Analyzer, RTCA (Multiple Plate) MP Station and RTCA Control Unit</td></tr><tr><td>Trypsin-EDTA (0.5%)</td><td>Gibco</td><td>15400-054&amp;nbsp;</td><td></td></tr></tbody></table>

using cell-substrate impedance and live cell imaging to measure real-time changes in cellular adhesion and de-adhesion induced by matrix modification

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and other matrix-modifying oxidants/enzymes released during inflammation) are implicated in triggering pathological changes in cellular function, phenotype and viability in a number of diseases. Here, we describe how cell-substrate impedance and live cell imaging approaches can be readily employed to accurately quantify real-time changes in cell adhesion and de-adhesion induced by matrix modification (using endothelial cells and myeloperoxidase as a pathophysiological matrix-modifying stimulus) with high temporal resolution and in a non-invasive manner. The xCELLigence cell-substrate impedance system continuously quantifies the area of cell-matrix adhesion by measuring the electrical impedance at the cell-substrate interface in cells grown on gold microelectrode arrays. Image analysis of time-lapse differential interference contrast movies quantifies changes in the projected area of individual cells over time, representing changes in the area of cell-matrix contact. Both techniques accurately quantify rapid changes to cellular adhesion and de-adhesion processes. Cell-substrate impedance on microelectrode biosensor arrays provides a platform for robust, high-throughput measurements. Live cell imaging analyses provide additional detail regarding the nature and dynamics of the morphological changes quantified by cell-substrate impedance measurements. These complementary approaches provide valuable new insights into how myeloperoxidase-catalyzed oxidative modification of subcellular extracellular matrix components triggers rapid changes in cell adhesion, morphology and signaling in endothelial cells. These approaches are also applicable for studying cellular adhesion dynamics in response to other matrix-modifying stimuli and in related adherent cells (e.g., epithelial cells).

Real-time quantification of endothelial cell de-adhesion from fibronectin in response to MPO-mediated fibronectin oxidation (Experiment 2). The seeding of endothelial cell suspensions onto native (MPO free) fibronectin or MPO-bearing fibronectin results in maximal cell attachment and spreading within 2 hr, as judged by a plateauing of cell index values in the cell substrate impedance measurements (Figure 3A). This initial phase of cell attachment and spreading is markedly reduced when MPO-mediated fibronectin oxidation is initiated prior to cell seeding in experiments performed according to the protocol detailed in Experiment 1 (data not shown; For details see9). Once maximal cell adhesion is established on native or MPO-bearing fibronectin, targeted fibronectin oxidation mediated by MPO-catalyzed HOCl (initiated by the addition of MPO&#8217;s co-substrate H2O2) causes a rapid decrease in the surface area of cell-matrix contact measured by cell-substrate impedance (Figure 3A,B) and by live cell imaging (Figure 3C; for a representative DIC movie, see Movie 1; cell index or cell area losses after H2O2 addition are minimal in the absence of MPO9). Whilst cell index and cell area changes during MPO-induced de-adhesion were very similar, the comparatively slower decreases in cell index values may reflect the insulating effects of cell membrane materials present in &#8216;cell-free&#8217; regions at the cell periphery during de-adhesion, which are not quantified in the projected cell area measurements. The rapid cellular de-adhesion apparent in endothelial cells bound to MPO-bearing fibronectin in response to H2O2 treatment is absent in cells treated with H2O2 alone (i.e., cells that do not contain MPO) (Figure 3A) and is blocked by MPO enzyme inhibitors or HOCl scavengers (data not shown; see9 for details), identifying that this process is dependent on the MPO-catalyzed production of HOCl (cf. Figure 1). Inhibition of myosin II motor function with blebbistatin inhibits the rate of endothelial cell de-adhesion measured by cell-substrate impedance (Figure 3B) and by live cell imaging (Figure 3C, Movie 2), identifying that cellular de-adhesion and contraction in response to MPO-catalyzed subcellular matrix oxidation is driven by actomyosin tensile forces9.
<img alt="Figure 1" src="/files/ftp_upload/52423/52423fig1highres.jpg" /> 
Figure 1. MPO-mediated subcellular matrix oxidation triggers cell-matrix de-adhesion and cell signaling. MPO triggers rapid de-adhesion responses and changes in adhesion-dependent signaling by mediating targeted oxidation of adhesive subendothelial matrix proteins, involving the following events9: (A) MPO binds avidly to the subendothelial matrix and uses H2O2 to generate the highly reactive oxidant HOCl that reacts locally with matrix proteins (e.g., fibronectin) and disrupts their cell adhesive properties. (B) This damage disrupts adhesive contacts at the cell-matrix interface, leading to (C) membrane retraction driven by unopposed tension in the actin cytoskeleton and the alteration of adhesion-dependent cell signaling pathways (Reproduced with permission from Rees et al.9). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52423/52423fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52423/52423fig2highres.jpg" /> 
Figure 2. Work-flow and principle of cell-substrate impedance analyses to quantify endothelial cell de-adhesion from fibronectin in response to MPO-mediated fibronectin oxidation (Experiment 2). Cell index values measured by the cell-substrate microarray impedance biosensor system reflect the ability of the cell membrane to limit field-induced ion currents at the electrode surface and are proportional to the area of cell&#8211;substrate contact10. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52423/52423fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52423/52423fig3highres.jpg" /> 
Figure 3. Real-time quantification of endothelial cell de-adhesion from fibronectin in response to MPO-mediated fibronectin oxidation (Experiment 2). Endothelial cell suspensions (in serum-free Medium 199 containing 1% BSA) were seeded onto native or MPO-bearing fibronectin (fibronectin coated at 5 g/ml, then incubated in the absence or presence of 20 nM MPO) in 96 well microelectrode arrays (cell-substrate impedance measurements) or 35 mm glass-bottom cell culture dishes (live cell imaging analyses). Cells were then incubated at 37&#176;C for 2 hr to allow maximal cell attachment and spreading before equilibrating cells with HBSS and initiating MPO-mediated fibronectin oxidation by adding H2O2 (10 &#956;M) (time of H2O2 addition set at t = 0 min). (A) Full time-course of cell-substrate impedance measurements before and after treatment with H2O2 in the presence (+MPO) and absence (-MPO) of MPO. (B) Cell-substrate impedance measurements and (C) cell area measurements by live cell imaging analyses, after treatment of MPO-containing cells with H2O2 in the presence (+ blebbistatin) and absence (-blebbistatin) of the myosin II inhibitor blebbistatin (40 &#956;M). Cell index and cell area values are normalized to values immediately prior to the time of H2O2 addition, which were given a value of 1. Cell index data represent the mean SEM, n = 6 replicate measurements from a representative experiment. Cell area values represent the mean SEM, n = 10 cells (2 replicate movies, 5 randomly selected cells per movie: see Movies 1 and 2). (Figure 3B, Figure 3C, Movie 1 and Movie 2 are reproduced with permission from Rees et al.9). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52423/52423fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52423/Movie 1.mov">Movie 1</a>.&#160;Time lapse DIC microscopy movie of endothelial cells adhered on MPO-bearing fibronectin over 0 &#8211; 15 min following exposure to H2O2 (10 &#956;M); 1 sec = 3 min.
<a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52423/Movie 2.mov">Movie 2</a>.&#160;Time lapse DIC microscopy movie of endothelial cells adhered on MPO-bearing fibronectin over 0 &#8211; 15 min following exposure to H2O2 (10 &#956;M) in the presence of blebbistatin (40 &#956;M); 1 sec = 3 min.

Watch this Scientific Journal Video about Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification at JoVE.

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and ...

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Universidad San Sebastian

Research

JoVE Journal

Bioengineering

10.9K Views.  University of New South Wales. Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Video: Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Preparation of Complaint Matrices for Quantifying Cellular Contraction

The regulation of cellular adhesion to the extracellular matrix (ECM) is essential for cell migration and ECM remodeling. Focal adhesions are macromolecular assemblies that couple the contractile F-actin cytoskeleton to the ECM. This connection allows for the transmission of intracellular mechanical forces across the cell membrane to the underlying substrate. Recent work has shown the mechanical properties of the ECM regulate focal adhesion and F-actin morphology as well as numerous physiological processes, including cell differentiation, division, proliferation and migration. Thus, the use of cell culture substrates has become an increasingly prevalent method to precisely control and modulate ECM mechanical properties.
To quantify traction forces at focal adhesions in an adherent cell, compliant substrates are used in conjunction with high-resolution imaging and computational techniques in a method termed traction force microscopy (TFM). This technique relies on measurements of the local magnitude and direction of substrate deformations induced by cellular contraction. In combination with high-resolution fluorescence microscopy of fluorescently tagged proteins, it is possible to correlate cytoskeletal organization and remodeling with traction forces.
Here we present a detailed experimental protocol for the preparation of two-dimensional, compliant matrices for the purpose of creating a cell culture substrate with a well-characterized, tunable mechanical stiffness, which is suitable for measuring cellular contraction. These protocols include the fabrication of polyacrylamide hydrogels, coating of ECM proteins on such gels, plating cells on gels, and high-resolution confocal microscopy using a perfusion chamber. Additionally, we provide a representative sample of data demonstrating location and magnitude of cellular forces using cited TFM protocols.

In this video, we demonstrate the experimental techniques used to fabricate compliant, extracellular matrix (ECM) coated substrates suitable for cell culture, and which are amenable to traction force microscopy and observing effects of ECM stiffness on cell behavior.

The regulation of cellular adhesion to the extracellular matrix (ECM) is essential for cell migration and ECM remodeling.  Focal adhesions are ...

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 ...

Electric Cell-substrate Impedance Sensing for the Quantification of Endothelial Proliferation, Barrier Function, and Motility

Electric Cell-substrate Impedance Sensing (ECIS) is an in vitro impedance measuring system to quantify the behavior of cells within adherent cell layers. To this end, cells are grown in special culture chambers on top of opposing, circular gold electrodes. A constant small alternating current is applied between the electrodes and the potential across is measured. The insulating properties of the cell membrane create a resistance towards the electrical current flow resulting in an increased electrical potential between the electrodes. Measuring cellular impedance in this manner allows the automated study of cell attachment, growth, morphology, function, and motility. Although the ECIS measurement itself is straightforward and easy to learn, the underlying theory is complex and selection of the right settings and correct analysis and interpretation of the data is not self-evident. Yet, a clear protocol describing the individual steps from the experimental design to preparation, realization, and analysis of the experiment is not available. In this article the basic measurement principle as well as possible applications, experimental considerations, advantages and limitations of the ECIS system are discussed. A guide is provided for the study of cell attachment, spreading and proliferation; quantification of cell behavior in a confluent layer, with regard to barrier function, cell motility, quality of cell-cell and cell-substrate adhesions; and quantification of wound healing and cellular responses to vasoactive stimuli. Representative results are discussed based on human microvascular (MVEC) and human umbilical vein endothelial cells (HUVEC), but are applicable to all adherent growing cells.

This protocol reviews Electric Cell-substrate Impedance Sensing, a method to record and analyze the impedance spectrum of adherent cells for the quantification of cell attachment, proliferation, motility, and cellular responses to pharmacological and toxic stimuli. Detection of endothelial permeability and assessment of cell-cell and cell-substrate contacts are emphasized.

Electric Cell-substrate Impedance Sensing (ECIS) is an in vitro impedance measuring system to quantify the behavior of cells within adherent cell layers. ...

Impedance-based Real-time Measurement of Cancer Cell Migration and Invasion

Cancer arises due to uncontrolled proliferation of cells initiated by genetic instability, mutations, and environmental and other stress factors. These acquired abnormalities in complex, multilayered molecular signaling networks induce aberrant cell proliferation and survival, extracellular matrix degradation, and metastasis to distant organs. Approximately 90% of cancer-related deaths are estimated to be caused by the direct or indirect effects of metastatic dissemination. Therefore, it is important to establish a highly reliable, comprehensive system to characterize cancer cell behaviors upon genetic and environmental manipulations. Such a system can give a clear understanding of the molecular regulation of cancer metastasis and the opportunity for successful development of stratified, precise therapeutic strategies. Hence, accurate determination of cancer cell behaviors such as migration and invasion with gain or loss of function of gene(s) allows assessment of the aggressive nature of cancer cells. The real-time measurement system based on cell impedance enables researchers to continually acquire data during a whole experiment and instantly compare and quantify the results under various experimental conditions. Unlike conventional methods, this method does not require fixation, staining, and sample processing to analyze cells that migrate or invade. This method paper emphasizes detailed procedures for real-time determination of migration and invasion of glioblastoma cancer cells.

Cancer is a lethal disease due to its ability to metastasize to different organs. Determining the ability of cancer cells to migrate and invade under various treatment conditions is crucial to assessing therapeutic strategies. This protocol presents a method to assess the real-time metastatic abilities of a glioblastoma cancer cell line.&#160;

Cancer arises due to uncontrolled proliferation of cells initiated by genetic instability, mutations, and environmental and other stress factors. These ...

Cancer Research

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic development, wound healing, and immune responses. However, cell migration is also a key mechanism in cancer enabling these cancer cells to detach from the primary tumor to start metastatic spreading. Within the past years various cell migration assays have been developed to analyze the migratory behavior of different cell types. Because the locomotory behavior of cells markedly differs between a two-dimensional (2D) and three-dimensional (3D) environment it can be assumed that the analysis of the migration of cells that are embedded within a 3D environment would yield in more significant cell migration data. The advantage of the described 3D collagen matrix migration assay is that cells are embedded within a physiological 3D network of collagen fibers representing the major component of the extracellular matrix. Due to time-lapse video microscopy real cell migration is measured allowing the determination of several migration parameters as well as their alterations in response to pro-migratory factors or inhibitors. Various cell types could be analyzed using this technique, including lymphocytes/leukocytes, stem cells, and tumor cells. Likewise, also cell clusters or spheroids could be embedded within the collagen matrix concomitant with analysis of the emigration of single cells from the cell cluster/ spheroid into the collagen lattice. We conclude that the 3D collagen matrix migration assay is a versatile method to analyze the migration of cells within a physiological-like 3D environment.

Cell migration is a biological phenomenon that is involved in a plethora of physiological, such as wound healing and immune responses, and pathophysiological processes, like cancer. The 3D-collagen matrix migration assay is a versatile tool to analyze the migratory properties of different cell types within in a 3D physiological-like environment.

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic ...

Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to tumor and cancer metastasis. Despite intense research efforts, the basic biochemical and biophysical principles of cell migration are still not fully understood, especially in the physiologically relevant three-dimensional (3D) microenvironments. Here, we describe an in vitro assay designed to allow quantitative examination of 3D cell migration behaviors. The method exploits the cell&#8217;s mechanosensing ability and propensity to migrate into previously unoccupied extracellular matrix (ECM). We use the invasion of highly invasive breast cancer cells, MDA-MB-231, in collagen gels as a model system. The spread of cell population and the migration dynamics of individual cells over weeks of culture can be monitored using live-cell imaging and analyzed to extract spatiotemporally-resolved data. Furthermore, the method is easily adaptable for diverse extracellular matrices, thus offering a simple yet powerful way to investigate the role of biophysical factors in the microenvironment on cell migration.

The mechanical properties and microstructure of the extracellular matrix strongly affect 3D migration of cells. An in vitro method to study the spatiotemporal cell migration behavior in biophysically variable environments, at both population and individual cell levels, is described.

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to ...

Real-Time Detection and Capture of Invasive Cell Subpopulations from Co-Cultures

Invasion and metastatic spread of cancer cells are the major cause of death from cancer. Assays developed early on to measure the invasive potential of cancer cell populations typically generate a single endpoint measurement that does not distinguish between cancer cell subpopulations with different invasive potential. Also, the tumor microenvironment consists of different resident stromal and immune cells that alter and participate in the invasive behavior of cancer cells. Invasion into tissues also plays a role in immune cell subpopulations fending off microorganisms or eliminating diseased cells from the parenchyma and endothelial cells during tissue remodeling and angiogenesis. Real-Time Cellular Analysis (RTCA) that utilizes impedance biosensors to monitor cell invasion was a major step forward beyond endpoint measurement of invasion: this provides continuous measurements over time and thus can reveal differences in invasion rates that are lost in the endpoint assay. Using current RTCA technology, we expanded dual-chamber arrays by adding a further chamber that can contain stromal and/or immune cells and allows measuring the rate of invasion under the influence of secreted factors from co-cultured stromal or immune cells over time. Beyond this, the unique design allows for detaching chambers at any time and isolating of the most invasive cancer cell, or other cell subpopulations that are present in heterogeneous mixes of tumor isolates tested. These most invasive cancer cells and other cell subpopulations drive malignant progression to metastatic disease, and their molecular characteristics are important for in-depth mechanistic studies, the development of diagnostic probes for their detection, and the assessment of vulnerabilities. Thus, the inclusion of small- or large-molecule drugs can be used to test the potential of therapies that target cancer and/or stromal cell subpopulations with the goal of inhibiting (e.g., cancer cells) or enhancing (e.g., immune cells) invasive behavior.

We describe an approach to detect and capture invasive cell subpopulations in real-time. The experimental design uses Real-Time Cellular Analysis by monitoring changes in the electric impedance of cells. Invasive cancer, immune, endothelial or stromal cells in complex tissues can be captured, and the impact of co-cultures can be assessed.

Invasion and metastatic spread of cancer cells are the major cause of death from cancer. Assays developed early on to measure the invasive potential of ...

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 ...

Biology

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts 

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Summary

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Chapters in this video

Preparation of Complaint Matrices for Quantifying Cellular Contraction

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

Electric Cell-substrate Impedance Sensing for the Quantification of Endothelial Proliferation, Barrier Function, and Motility

Impedance-based Real-time Measurement of Cancer Cell Migration and Invasion

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

Real-Time Detection and Capture of Invasive Cell Subpopulations from Co-Cultures

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

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts