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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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 ...

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 border="0" cellpadding="0" cellspacing="0" width="1328">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Name of Material/ Equipment</td>
			<td style="width:223px;">Company</td>
			<td style="width:151px;">Catalog Number</td>
			<td style="width:551px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">96 well gold cell-substrate impedance microelectrode array</td>
			<td style="width:223px;">ACEA Biosciences / Roche</td>
			<td style="width:151px;">E-Plate 96</td>
			<td>single-use plate used for performing cell-based assays on the xCELLigence system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">blebbistatin</td>
			<td style="width:223px;">Sigma</td>
			<td style="width:151px;">B0560</td>
			<td>selective inhibitor of non-muscle myosin-II</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Bovine Aortic Endothelial Cells, cryopreserved&#160;</td>
			<td style="width:223px;">Lonza</td>
			<td style="width:151px;">BW-6001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Bovine serum albumin</td>
			<td style="width:223px;">Sigma</td>
			<td style="width:151px;">05470</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">EGM-2 BulletKit</td>
			<td style="width:223px;">Lonza</td>
			<td style="width:151px;">CC-3162</td>
			<td>endothelial cell growth media kit</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Fibronectin, lyophilized powder</td>
			<td style="width:223px;">Sigma</td>
			<td style="width:151px;">F-4759</td>
			<td>from bovine plasma</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Fluorodish 35 mm glass-bottomed cell culture dish</td>
			<td style="width:223px;">World Precision Instruments</td>
			<td style="width:151px;">FD35-100</td>
			<td>Cell cuture dish with optical quality glass bottom for imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Gelatin from bovine skin</td>
			<td style="width:223px;">Sigma</td>
			<td style="width:151px;">G9391</td>
			<td>cell culture substratum</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Hank&#39;s Balanced Salt Solution</td>
			<td style="width:223px;">Life Technologies</td>
			<td style="width:151px;">14025076</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Hydrogen peroxide</td>
			<td style="width:223px;">Merck</td>
			<td style="width:151px;">107298</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Medium-199</td>
			<td style="width:223px;">Life Technologies</td>
			<td style="width:151px;">11150-059</td>
			<td>serum-free cell media</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Methionine</td>
			<td style="width:223px;">Sigma</td>
			<td style="width:151px;">M9500</td>
			<td>quenches chlorinating oxidants generated by myeloperoxidase</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Myeloperoxidase, Human Polymorphonuclear Leukocytes</td>
			<td style="width:223px;">Millipore</td>
			<td style="width:151px;">475911</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:405px;">RTCA MP Instrument / xCELLigence cell substrate impedance system</td>
			<td style="width:223px;">ACEA Biosciences / Roche</td>
			<td style="width:151px;">-</td>
			<td>Consists of RTCA Analyzer, RTCA (Multiple Plate) MP Station and RTCA Control Unit</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:405px;">Trypsin-EDTA (0.5%)</td>
			<td>Gibco</td>
			<td>15400-054&#160;</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 MP...

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 ...

using-cell-substrate-impedance-live-cell-imaging-to-measure-real-time

Research

JoVE Journal

Bioengineering

10.9K 조회수.  University of New South Wales.  여기서, 우리는 지속적으로 셀 기판 임피던스와 생균 영상 분석에 의해 비 침습성 방식으로 높은 시간 해상도를 가진 세포 부착 및 디 - 접착 공정을 정량화하는 프로토콜을 제시한다. 이러한 접근은 매트릭스 수정 및 접착 의존성 시그널링 이벤트들이 시간적인 관계에 의해 트리거 세포 접착 / 디 접착 공정의 역학을 보여준다. 

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

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 ...

Harvesting Murine Alveolar Macrophages and Evaluating Cellular Activation Induced by Polyanhydride Nanoparticles

Biodegradable nanoparticles have emerged as a versatile platform for the design and implementation of new intranasal vaccines against respiratory infectious diseases. Specifically, polyanhydride nanoparticles composed of the aliphatic sebacic acid (SA), the aromatic 1,6-bis(p-carboxyphenoxy)hexane (CPH), or the amphiphilic 1,8-bis(p-carboxyphenoxy)-3,6-dioxaoctane (CPTEG) display unique bulk and surface erosion kinetics1,2 and can be exploited to slowly release functional biomolecules (e.g., protein antigens, immunoglobulins, etc.) in vivo3,4,5. These nanoparticles also possess intrinsic adjuvant activity, making them an excellent choice for a vaccine delivery platform6,7,8.
	In order to elucidate the mechanisms governing the activation of innate immunity following intranasal mucosal vaccination, one must evaluate the molecular and cellular responses of the antigen presenting cells (APCs) responsible for initiating immune responses. Dendritic cells are the principal APCs found in conducting airways, while alveolar macrophages (AM&#632;) predominate in the lung parenchyma9,10,11. AM&#632; are highly efficient in clearing the lungs of microbial pathogens and cell debris12,13. In addition, this cell type plays a valuable role in the transport of microbial antigens to the draining lymph nodes, which is an important first step in the initiation of an adaptive immune response9. AM&#632; also express elevated levels of innate pattern recognition and scavenger receptors, secrete pro-inflammatory mediators, and prime na&iuml;ve T cells12,14. A relatively pure population of AM&#632; (e.g., greater than 80%) can easily be obtained via lung lavage for study in the laboratory. Resident AM&#632; harvested from immune competent animals provide a representative phenotype of the macrophages that will encounter the particle-based vaccine in vivo. Herein, we describe the protocols used to harvest and culture AM&#632; from mice and examine the activation phenotype of the macrophages following treatment with polyanhydride nanoparticles in vitro.

Herein, we describe protocols for harvesting murine alveolar macrophages, which are resident innate immune cells in the lung, and examining their activation in response to co-culture with polyanhydride nanoparticles.

Biodegradable nanoparticles have emerged as a  versatile platform for the design and implementation of new intranasal vaccines  against respiratory ...

Simultaneously Capturing Real-time Images in Two Emission Channels Using a Dual Camera Emission Splitting System: Applications to Cell Adhesion

Multi-color immunofluorescence microscopy to detect specific molecules in the cell membrane can be coupled with parallel plate flow chamber assays to investigate mechanisms governing cell adhesion under dynamic flow conditions. For instance, cancer cells labeled with multiple fluorophores can be perfused over a potentially reactive substrate to model mechanisms of cancer metastasis. However, multi-channel single camera systems and color cameras exhibit shortcomings in image acquisition for real-time live cell analysis. To overcome these limitations, we used a dual camera emission splitting system to simultaneously capture real-time image sequences of fluorescently labeled cells in the flow chamber. Dual camera emission splitting systems filter defined wavelength ranges into two monochrome CCD cameras, thereby simultaneously capturing two spatially identical but fluorophore-specific images. Subsequently, psuedocolored one-channel images are combined into a single real-time merged sequence that can reveal multiple target molecules on cells moving rapidly across a region of interest.

Dual camera emission splitting systems for two-color fluorescence microscopy generate real-time image sequences with exceptional optical and temporal resolution, a requirement of certain live cell assays including parallel plate flow chamber adhesion assays. When software is employed to merge images from simultaneously acquired emission channels, pseudocolored image sequences are produced.

Multi-color immunofluorescence microscopy to detect specific molecules in the cell membrane can be coupled with parallel plate flow chamber assays to ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

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. ...

Universal Hand-held Three-dimensional Optoacoustic Imaging Probe for Deep Tissue Human Angiography and Functional Preclinical Studies in Real Time

The exclusive combination of high optical contrast and excellent spatial resolution makes optoacoustics (photoacoustics) ideal for simultaneously attaining anatomical, functional and molecular contrast in deep optically opaque tissues. While enormous potential has been recently demonstrated in the application of optoacoustics for small animal research, vast efforts have also been undertaken in translating this imaging technology into clinical practice. We present here a newly developed optoacoustic tomography approach capable of delivering high resolution and spectrally enriched volumetric images of tissue morphology and function in real time. A detailed description of the experimental protocol for operating with the imaging system in both hand-held and stationary modes is provided and showcased for different potential scenarios involving functional and molecular studies in murine models and humans. The possibility for real time visualization in three dimensions along with the versatile handheld design of the imaging probe make the newly developed approach unique among the pantheon of imaging modalities used in today&#8217;s preclinical research and clinical practice.

We provide herein a detailed description of the experimental protocol for imaging with a newly developed hand-held optoacoustic (photoacoustic) system for three-dimensional functional and molecular imaging in real time. The demonstrated powerful performance and versatility may define new application areas of the optoacoustic technology in preclinical research and clinical practice.

The exclusive combination of high optical contrast and excellent spatial resolution makes optoacoustics (photoacoustics) ideal for simultaneously ...

Hand-held Clinical Photoacoustic Imaging System for Real-time Non-invasive Small Animal Imaging

Translation of photoacoustic imaging into the clinic is a major challenge. Handheld real-time clinical photoacoustic imaging systems are very rare. Here, we report a combined photoacoustic and clinical ultrasound imaging system by integrating an ultrasound probe with light delivery for small animal imaging. We demonstrate this by showing sentinel lymph node imaging in small animals along with minimally invasive real-time needle guidance. A clinical ultrasound platform with access to raw channel data allows the integration of photoacoustic imaging leading to a handheld real-time clinical photoacoustic imaging system. Methylene blue was used for sentinel lymph node imaging at 675 nm wavelength. Additionally, needle guidance with dual modal ultrasound and photoacoustic imaging was shown using the imaging system. Depth imaging of up to 1.5 cm was demonstrated with a 10 Hz laser at a photoacoustic imaging frame rate of 5 frames per second.

A clinical handheld photoacoustic imaging system will be demonstrated for real-time non-invasive small animal imaging.

Translation of photoacoustic imaging into the clinic is a major challenge. Handheld real-time clinical photoacoustic imaging systems are very rare. Here, ...

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 ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of cellular function. The ECM plays a particularly important role in adipose tissue function in obesity, and alterations in adipose tissue ECM deposition and composition are associated with metabolic disease in mice and humans. Tractable in vitro models that permit dissection of the roles of the ECM and cells in contributing to global tissue phenotype are sparse. We describe a novel 3D in vitro model of human ECM-adipocyte culture that permits study of the specific roles of the ECM and adipocytes in regulating adipose tissue metabolic phenotype. Human adipose tissue is decellularized to isolate ECM, which is subsequently repopulated with preadipocytes that are then differentiated within the ECM into mature adipocytes. This method creates ECM-adipocyte constructs that are metabolically active and retain characteristics of the tissues and patients from which they are derived. We have used this system to demonstrate disease-specific ECM-adipocyte crosstalk in human adipose tissue. This culture model provides a tool for dissecting the roles of the ECM and adipocytes in contributing to global adipose tissue metabolic phenotype and permits study of the role of the ECM in regulating adipose tissue homeostasis.

We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of ...