We are most grateful to Dr. Belinda Willard, Proteomics and Metabolomics Laboratory, Lerner Research Institute, Cleveland Clinic, for conducting the proteomics analysis of RCS ECM. We acknowledge the financial support of theMedical Research Council UK, grant number K018043, to JCA.

1. Removal of Cells with Ammonium Hydroxide Solution
<ol>
	<li>Prepare Adherent Cells by Plating at the Appropriate Density. 
	NOTE: The cells can be any adherent cell type that produces sufficient ECM for analysis. Here, we describe the use of COS-7 cells, an African green monkey kidney fibroblast-like cell line that contains SV-40 viral DNA sequences; RCS, a rat chondrosarcoma cell line; or normal human dermal fibroblast (HDF) strains from juvenile foreskin. HDF are used from passage 1 to passage 8 only.
	<ol>
		<li>Plate the cells on coverslips for the imaging of live cells and ECM, for fluorescence microscopy studies of fixed ECM, or for preparation of cell-derived ECM for small-scale functional assays. Plate the cells on cell culture dishes for SDS-PAGE analysis, immunoblot, or proteomics studies. 
		NOTE: The cell culture conditions (number of cells to plate and culture medium) will depend on the cell type. The cell number to plate will need to be established empirically for the particular cell line or cell strain.</li>
		<li>Allow a suitable time for the cells to deposit ECM, typically &#62;16 h. NOTE: The time period will depend on the cell type and will need to be determined empirically. If conducting ectopic expression, allow appropriate time for the expression of the transfected protein of interest and for the deposition of ECM.</li>
	</ol>
	</li>
	<li>In an Extractor Hood, Prepare 20 mM Ammonium Hydroxide in a Suitable Vessel by Diluting the Stock Solution 1/14 with De-ionized H2O.
	<ol>
		<li>Remove the cells from the incubator and gently remove culture medium. Add phosphate-buffered saline (PBS) without Ca2+/Mg2+ by gently pouring against the walls of the dish. Rock the dish twice and remove the liquid with a plastic transfer pipette. Repeat twice more.</li>
	</ol>
	</li>
	<li>In an extractor hood, tilt each dish and remove the PBS from step 1.2 with a plastic transfer pipette. Add 3 mL of ammonium hydroxide per 100-mm dish and incubate them at room temperature for 5 min. During the 5-min incubation period, gently agitate the dish every min to ensure the lysis of all the cells. Steps 1.4-1.7 will also be carried out in the extractor hood. 
	NOTE: Alternative cell removal reagents include 2 M or 8 M urea, which are incubated with the cells for 10 min.</li>
	<li>Add copious amounts of de-ionized H2O to each dish, at least 20 mL per 100 mm dish, with rocking. Dispose of the ammonium hydroxide-solubilized material-which is composed of ammonium hydroxide, lysed cells, and de-ionized H2O-by aspiration with a transfer pipette. Transfer this waste solution into a container for liquid waste.</li>
	<li>Wash the insoluble ECM layer in copious de-ionized H2O four more times to ensure the complete removal of all the ammonium hydroxide-solubilized material.</li>
	<li>For examination of the ECM by immunofluorescence, fix the ECM by adding 2% paraformaldehyde (PFA; 3 mL per 100 mm dish) at room temperature for 10 min. 
	Caution: PFA is very hazardous in case of skin or eye contact and severe overexposure can produce lung damage, choking, unconsciousness, or death. It should only be handled in an extractor hood, and personal protective equipment should be worn.</li>
	<li>Wash each fixed sample twice with PBS, as in step 1.2, and dispose of the PFA solution into a suitable liquid waste container. If necessary, store the ECM samples in PBS at 4 &#176;C before preparing them for fluorescence microscopy.</li>
</ol>
2. Tracking Deposition of an ECM Protein by Confocal Microscopy Imaging of Live Cells
<ol>
	<li>Count the cells under a hemocytometer. For COS-7 cells, plate 250,000 cells onto a 60 mm cell culture dish for transfection. 
	NOTE: For live cell imaging, the cells must be transfected with a vector encoding the ECM protein of interest, fused in frame with a genetically-encoded fluorescent tag. This is to enable the identification of the ECM protein of interest during live-cell imaging.</li>
	<li>After a suitable time for protein expression (e.g., 16 h) remove the cells from the dish with trypsin-EDTA solution, count them in a hemocytometer, and plate ~150,000 cells onto a 35-mm gridded, glass-bottomed dish for imaging. Allow 2 h for the cells to reattach and to begin ECM deposition (the time period will depend on the cell type and must be established empirically). 
	NOTE: Use a cell medium that does not contain phenol red, as this can give a red tinge that affects image quality.</li>
	<li>During the above 2 h period, set up a confocal microscope with a 37 &#176;C incubator chamber and a CO2 supply. Allow the temperature in the chamber and in the dish to stabilize to 37 &#176;C before imaging; this can take up to 1 h.</li>
	<li>Incubate the cells plated onto the 35 mm gridded dish with a fluorescent cell marker for 30 min. Then, wash the cells twice in phenol red-free culture medium prior to imaging. 
	NOTE: A cytoplasmic fluorescent marker can be used to visualize cell volumes, or a membrane-incorporating marker can be used to visualize cell edges.</li>
	<li>Using the 20X objective and phase contrast on a confocal microscope, identify an appropriate grid square that contains a number (&#62;3) of adherent, healthy cells. Confirm the expression of the target protein of choice in the cells within the selected grid square using the 63X or 100X objectives and the appropriate wavelengths under fluorescence microscopy.</li>
	<li>Set up fluorescence and phase-contrast time-lapse imaging using a z-stack software. Set the &#34;Begin&#34; stack to be just below the ECM layer and the &#34;End&#34; stack to be just above the cell body. Set the z-step size to 0.3 &#181;m and the z-volume to between a 5 and 10 &#181;m depth in total, depending on the cell type. This will give between 15-30 z-sections per time point.</li>
	<li>Capture fluorescence (for red fluorescent protein (RFP), excitation = 555 nm and emission = 584 nm) and phase-contrast images periodically over a set time period. For example, use the time-lapse software to set the time interval at 2 min and the duration to 2 h. Select the &#34;start&#34; button to begin capture of z-section images over the defined time period.</li>
	<li>At the end of the time period, remove the cells from dish, as described in steps 1.1-1.5, to confirm the localization of the fluorescently-tagged protein of interest in the ECM.</li>
	<li>Use phase-contrast imaging and the 20X objective to relocate the relevant grid square.</li>
	<li>Switch to the 63X objective and identify the ECM layer under fluorescence microscopy. Compare this image to the pre-extraction image</li>
</ol>
3. Sterile Preparation of Cell-derived ECM for Cell Functional Assays
<ol>
	<li>Grow one population of cells (the &#34;matrix producer&#34; cells) on coverslips and another population (the &#34;test&#34; cells) in standard culture in a 100 mm cell culture dish for at least 24 h. 
	NOTE: These can be any two different adherent cell types, and the experimental design can include transfection of one or both cell populations to express a fluorescently-tagged ECM protein.</li>
	<li>In a laminar flow hood, as used for cell culture, prepare sterile solutions of PBS without Ca2+/Mg2+, 20 mM ammonium hydroxide (see step 1.3), and de-ionized H2O by passing each through a sterile 0.2 &#181;m filter into an appropriate sterile container.</li>
	<li>Maintaining sterile technique, gently wash the &#34;matrix producer&#34; cells on coverslips three times with sterile PBS (see step 1.2).</li>
	<li>Remove the PBS by aspiration with a sterile pipette and dispose it in a suitable liquid waste container. Add 20 mM sterile ammonium hydroxide (3 mL/100 mm dish) and incubate them for 5 min with rocking at intervals.</li>
	<li>Add copious amounts of sterile de-ionized H2O to the &#34;matrix producer&#34; dish, at least 20 mL per 100 mm dish, with rocking, and pour away the cell lysate into a suitable waste container.</li>
	<li>Repeat step 3.5 four more times to ensure the complete removal of the ammonium hydroxide-solubilized material. 
	NOTE: The ECM deposited by the &#34;matrix producer&#34; cells on the coverslips then provides a sterile ECM for functional assays with any test cells of interest.</li>
	<li>Incubate the test cells from the cell stock dish with 1.5 mL of trypsin-EDTA solution per 100 mm dish (0.05% w/v trypsin and 0.02% w/v EDTA in Hank&#39;s balanced salt solution) until the test cells are detached from the dish. Add cell medium, centrifuge the test cells at 400 x g for 5 min, and remove the supernatant liquid.</li>
	<li>Resuspend the test cells in fresh, sterile medium and count them in a hemocytometer. Plate an appropriate number of these test cells onto the sterile, isolated ECM on coverslips. Culture them in the appropriate cell medium for 2-3 h, or for the time period required for the experimental design.</li>
	<li>To examine the organization of the actin cytoskeleton, fix the test cells in 2% PFA and stain them with fluorescein isothiocyanate (FITC)-phalloidin (excitation = 490 nm and emission = 525 nm) to visualize F-actin and with 4&#39;,6-diamidino-2-phenylindole (DAPI; excitation = 358 nm and emission = 461 nm) to visualize the nuclei11. 
	NOTE: Compare the morphology and actin organization in the test cells plated on heterologous cell-derived ECM with test cells plated on their own ECM.</li>
</ol>
4. Isolation of ECM for SDS-PAGE, Immunoblot Analysis, or Proteomics
<ol>
	<li>Grow adherent cells of interest on 100 mm cell culture dishes (5 to 10 dishes for proteomics or 1-2 dishes for immunoblotting) for 24-96 h, as appropriate for the cell type, to allow secretion and deposition of ECM.</li>
	<li>Remove the cells as described in steps 1.1-1.5.</li>
	<li>Tilt each plate at an angle to drain residual H2O to the bottom of the dish, and carefully remove any remaining H2O with a p200 pipet until the plate is completely dry.</li>
	<li>In parallel, heat SDS-PAGE sample buffer containing 100 mM dithiothreitol (DTT) to 95 &#176;C for 2 min using a heat block, and then add it to the plate. 200 &#181;L of sample buffer per 100 mm plate is appropriate for samples to be examined by immunoblot.
	<ol>
		<li>To analyze ECM by mass spectrometry, achieve a more concentrated sample by collecting the SDS-PAGE sample buffer from the dish (as described in steps 4.5 and 4.6, below), re-heating it to 95 &#176;C, and using the same aliquot across 2-3 additional plates.</li>
	</ol>
	</li>
	<li>Use a cell scraper to thoroughly scrape the ECM off the dish, ensuring that all areas of the dish have been scraped.</li>
	<li>With the cell scraper, gather the sample to one side of the dish and remove it with a 200 &#181;L pipette tip into a tube.</li>
	<li>Transfer any residual SDS-PAGE sample buffer extract from the cell scraper tip into the tube to minimize the loss of ECM material. For a concentrated sample, re-heat the same SDS-PAGE sample buffer aliquot to 95 &#176;C and re-use it across 2-3 additional plates.</li>
	<li>Resolve the ECM proteins by SDS-PAGE12, using either 7% or 4-7% gradient polyacrylamide gels. 
	NOTE: It is convenient to use pre-stained protein markers.</li>
	<li>To increase the resolution of the high-molecular-weight ECM proteins, extend the gel running time such that the 37 kDa molecular weight marker runs close to the bottom of the gel and molecular weight markers &#60;37 kDa have run off the gel.</li>
	<li>For proteomic analysis of ECM proteins, stain the gel with a protein stain and capture an image. Cut bands of interest from the gel, digest them with trypsin, and analyze them by matrix-assisted laser desorption/ionization (MALDI)-mass spectrometry13.</li>
	<li>Alternatively, for a more comprehensive analysis of the ECM extract and to analyze the entire sample, slice the gel lane into sections, digest them with trypsin, and perform liquid chromatography-mass spectrometry (LC-MS)14.</li>
	<li>Alternatively, for immunoblotting15, transfer proteins from the SDS-PAGE gel onto polyvinylidene fluoride (PVDF) membrane at 15 V for at least 1 h 10 min (semi-dry method) to ensure the transfer of the high-molecular-weight ECM proteins. Visualize the total protein transferred to the membrane with a Ponceau S stain.</li>
	<li>Following step 4.11, use antibodies to establish the presence and/or make a semi-quantitative analysis of individual ECM proteins15.
	<ol>
		<li>Block the membrane with 2% (w/v) milk in TBS-Tween 20 (blocking buffer) overnight at 4 &#176;C. Dilute specific antibodies to ECM proteins in blocking buffer and incubate with the membrane for 1.5 h at room temperature (antibody to fibronectin diluted 1/600 and antibody to thrombospondin-1 diluted 1/150; Supplementary Table 1).</li>
		<li>Test the quality of the ECM isolation by re-probing the same blot with antibodies to abundant intracellular proteins, such as actin or tubulin (anti-actin diluted 1/10,000 and anti-tubulin diluted 1/5,000).</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

Critical Steps within the Protocol
The method has some critical steps that must be followed to ensure the successful isolation and analysis of ECM. For example, ammonium hydroxide is used to remove the cells and to isolate ECM for analysis. Although ammonium hydroxide is stable in aqueous solutions in the dark and can be stored at room temperature, bottles must be tightly re-sealed between each use. Because the gas may evaporate from the solution, thus reducing the concentration, we strongly recommend starting a fresh bottle every 6-8 weeks. In addition, the working solution should be made freshly for each experiment. It is also essential that the cells are fully removed with copious washes in de-ionized water. If these steps are not performed adequately, cellular material may remain on the dish and contaminate downstream analyses. If cells have been grown for 10 days or longer, additional water washes may be needed. It is important to note that the isolated ECM layer is typically very thin in comparison to the cells11, so care is needed when identifying the ECM during imaging. For effective extraction of the isolated ECM into SDS-PAGE sample buffer, it is essential that the sample buffer contains a reducing agent. For the most effective removal of the ECM, boiling-hot sample buffer must be used. For experiments in which ECM samples will be resolved by SDS-PAGE electrophoresis for protein staining or proteomics, it is critical to achieve a concentrated sample by scraping several plates-worth of ECM into the same aliquot of sample buffer.
Modifications and Troubleshooting
The production and secretion of ECM proteins can vary significantly between cell types, so time periods for secretion and deposition of ECM should be determined de novo for each cell type. It is also important to be aware that ECM composition will change dynamically in relation to cell density and time in culture18. This factor should be taken into account when designing experiments. Clearly, the selection of a cell type for this method is important, particularly when extracting ECM for analysis by mass spectrometry, which may require a substantial amount of total ECM protein.
Limitations of the Technique
Cells that secrete very low levels of ECM proteins will be more challenging for use with this method. Non-adherent cells, 3D cell cultures, or cell invasion assays are unsuitable at present for ECM isolation by this method. The method is most suitable for use with standard &#34;2-dimensional&#34; cell cultures. Because the ammonium hydroxide extraction removes the cells completely, ECM associated with the upper sides of cells and secreted, but non-cross-linked, ECM proteins are also removed, leaving on the dish a thin layer of assembled ECM from below and around the bases of the cells11,17. It is complex to quantify how much ECM is released by the ammonium hydroxide extraction, because the released material also includes ECM proteins that are intracellular either prior to secretion or after uptake from the cell surface.
Significance of the Technique with Respect to Existing/Alternative Methods
The ability to conduct a wide range of experiments with isolated ECM is important in the context of cell biology and tissue physiology, and it also has implications for the fields of tissue regeneration and tissue engineering. The method has advantages over gels formed from purified collagens or other purified ECM proteins in that the cells assemble complex ECM structures at their native size, with native cross-linking mechanisms and with individual ECM proteins present in ratios appropriate to the cell type of origin. For example, the proteomic study of RCS ECM demonstrated abundant matrilins and COMP/TSP5, as expected for a cartilaginous ECM19,20 (Figure 4). In general, the analysis of cell-derived ECM is hampered by its nature as an extensive, multi-protein network that results in covalent crosslinking and the insolubility of many ECM proteins, and also by the potential contamination of ECM extracts with intracellular proteins. A multi-step procedure designed to isolate the ECM fraction from tissues for proteomic analysis yielded 8% of ECM proteins in the total set of proteins identified21. However, peptides from ECM proteins were 73% of the total peptides identified. This method for isolation and analysis of cell-derived ECM rapidly and reliably removes cellular material whilst retaining ECM proteins. This method can be used for a range of cell types and downstream applications and facilitates the analysis of ECM biology and cell-ECM interactions in cell culture. Our protocols detail how the method can be applied at different scales to address a wide range of experimental questions with regard to ECM organization, dynamics, or composition, and to the functional effects of isolated ECM on cells.
Future Applications or Directions after Mastering This Technique
Looking to the future, the method could be adapted for use with 3D cell cultures; this would expand its physiological relevance. Decellularized native tissue has great potential as a scaffold for the regeneration of lost or damaged tissue and as an alternative to transplantation, such as after heart failure22. It has the potential to overcome the problems of donor availability and immunorejection. Future applications of the method described in this report could investigate its use with 3D cell cultures or tissue decellularization, which would further increase the scope of this approach.

Over the last few decades, the extracellular matrix (ECM) has become recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and pathological processes. At the tissue level, the ECM influences cell signaling, motility, differentiation, angiogenesis, stem cell biology, tumorigenesis, fibrosis, etc.1,2. The study of ECM organization and ECM-dependent processes thus has wide implications for cell biology and tissue physiology. To reach a mechanistic understanding of ECM composition, organization, and functional properties, methods for the accurate isolation of ECM are required. Whereas the earliest identification of ECM proteins relied upon isolation from tissues3, the preparation of ECM from cultured cells has now become more prevalent.
The isolation and analysis of cell-derived ECM is complicated for two main reasons. Firstly, the presence of cells and their abundant intracellular proteins can make it difficult to isolate the ECM as a discrete extracellular structure. Indeed, some ECM proteins have roles inside the cell as well as in the ECM4; therefore, the efficient removal of intracellular proteins from cell-derived ECM is vital if the study of proteins within the ECM is not to be confused with their roles inside the cell. Secondly, cell-derived ECM is composed of many large, oligomeric proteins, which are often covalently cross-linked upon ECM assembly and are therefore insoluble in standard detergents. These properties can complicate extraction and the further analysis of ECM. To address these issues, a method is required that enables the efficient separation of ECM proteins from cellular components.
Several methods have been described in the literature for the isolation of ECM from either cell culture or tissue extracts. Many of these methods are aimed at the extraction of the abundant ECM protein, collagen, from tissues and include the use of neutral salts3, acidic conditions5,6, or pepsin7. Isolation of total ECM from tissue extracts often involves decellularization of the tissue prior to the isolation of the ECM. For example, a sequential extraction method has been described that isolates ECM from human cardiac tissue8. First, loosely-bound ECM proteins were extracted with 0.5 M NaCl before sodium dodecyl sulphate (SDS) was used to remove the cells. Finally, the remaining ECM proteins were extracted using 4 M guanidine8. ECM can also be solubilized from decellularized samples using 8 M urea9. Other techniques use detergents, such as deoxycholic acid10, to extract both cells and ECM before separating insoluble ECM proteins from the soluble cell lysate by centrifugation.
The method for isolation and analysis of cell-derived ECM described in this report provides a reproducible method for removing cell material, leaving cell-derived ECM that can be analyzed by in situ immunofluorescence or extracted for further biochemical analysis. This method can be adapted for any adherent cell type and can be scaled up for downstream procedures, such as immunoblotting or mass spectrometry, or for utilization of the isolated ECM in functional studies. The method can also be used in conjunction with confocal microscopy of live cells to track ECM deposition of a tagged protein of interest in real time. This is achieved through the use of a gridded, glass-bottomed dish. Overall, the approach provides an accurate isolation of cell-derived ECM and also the scope to identify and monitor the deposition and dynamics of individual ECM proteins.

<table><tbody><tr><td>Antibody to COL1A1</td><td>Novus</td><td>NB600-408</td><td>Rabbit polyclonal. Reacts with other fibrillar collagens as well as collagen I. IF: 1/200</td></tr><tr><td>Antibody to fibronectin</td><td>Sigma</td><td>F3648</td><td>Rabbit polyclonal. WB: 1/600 for 1.5 h. IF: 1/200</td></tr><tr><td>Antibody to thrombospondin 1</td><td>ThermoFisher&amp;nbsp;</td><td>MA5-13398</td><td>Mouse monoclonal clone A6.1. WB: 1/150 for 1.5 h</td></tr><tr><td>Antibody to RFP</td><td>Abcam</td><td>ab62341</td><td>Rabbit polyclonal. WB: 1/2,000 for 2 h</td></tr><tr><td>Antibody to &amp;beta;-actin</td><td>Sigma</td><td>A1978</td><td>Mouse monoclonal clone AC-15. WB: 1/10,000 for 1.5 h</td></tr><tr><td>Antibody to &amp;alpha;-tubulin</td><td>Sigma</td><td>T9026</td><td>Mouse monoclonal clone DM1A. WB: 1/5,000 for 1.5 h</td></tr><tr><td>Cell Tracker Green</td><td>ThermoFisher&amp;nbsp;</td><td>C2925</td><td>Fluorescent cell marker. Use at 1 &amp;micro;M for 30 min</td></tr><tr><td>Fluorescein isothiocyanate (FITC)-Phalloidin</td><td>Sigma</td><td>P-5282</td><td>Stock = 50 mg/mL in DMSO. 1/50 for 45 min</td></tr><tr><td>FITC-conjugated goat anti-mouse IgG</td><td>Sigma</td><td>F8771</td><td>IF: 1/50 for 1 h</td></tr><tr><td>FITC-conjugated sheep anti-rabbit IgG</td><td>SIgma</td><td>F7512</td><td>IF: 1/50 for 1 h</td></tr><tr><td>Horseradish peroxidase (HRP)-conjugated goat anti-mouse IgG</td><td>LI-COR</td><td>926-80010</td><td>WB: 1/50,000 for 1 h</td></tr><tr><td>HRP-conjugated goat anti-rabbit IgG</td><td>LI-COR</td><td>926-80011</td><td>WB: 1/100,000 for 1 h</td></tr><tr><td>Vectorshield mounting medium with DAPI</td><td>Vector</td><td>H-1200</td><td>Store at 4 &amp;deg;C in the dark</td></tr><tr><td>Dulbecco&#039;s Modified Eagle&#039;s Medium</td><td>Sigma</td><td>D6429</td><td>Warm before use</td></tr><tr><td>Fibroblast growth medium</td><td>PromoCell</td><td>C-23010</td><td>Warm before use, supplement with 50 &amp;micro;g/mL ascorbic acid</td></tr><tr><td>Phenol red-free DMEM</td><td>ThermoFisher</td><td>21063-029</td><td>Warm before use</td></tr><tr><td>Trypsin-EDTA solution</td><td>Sigma</td><td>T3924</td><td>Warm before use</td></tr><tr><td>Gridded glass-bottomed dish</td><td>MatTek</td><td>P35G-2-14-C-GRID</td><td></td></tr><tr><td>NH&lt;sub&gt;4&lt;/sub&gt;OH, 28-30% solution</td><td>Sigma</td><td>221228</td><td>Dilute to 20 mM in de-ionised water. Once opened store in the dark, tightly capped.</td></tr><tr><td>Paraformaldehyde, 16% solution</td><td>Alfa Aesar</td><td>43368</td><td>Dilute to 2% (v/v) in PBS</td></tr><tr><td>Deoxycholic acid</td><td>Sigma</td><td>D2510</td><td>Use at 2% (v/v) final concentration</td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>T8787</td><td>Dilute to 0.5% (v/v) final concentration</td></tr><tr><td>GelCode Blue stain reagent</td><td>ThermoFisher&amp;nbsp;</td><td>24590</td><td>Protein stain for SDS-PAGE gels</td></tr><tr><td>Precision Plus protein standards</td><td>Biorad</td><td>161-0374</td><td>Protein standards for SDS-PAGE gels</td></tr><tr><td>Trans-Blot SD Semi Dry transfer cell</td><td>Biorad</td><td>1703940</td><td>Efficient transfer of high molecular weight proteins</td></tr><tr><td>PVDF transfer membrane</td><td>Millipore</td><td>ISEQ00010</td><td>Immobilon-PSQ&amp;nbsp;</td></tr><tr><td>Ponceau S stain</td><td>Sigma</td><td>P7170</td><td>Reversible protein stain for PVDF membrane</td></tr><tr><td>WesternSure Enhanced chemiluminescence (ECL) substrate</td><td>LI-COR</td><td>926-80200</td><td></td></tr><tr><td>High performance chemiluminescence film</td><td>GE Healthcare</td><td>28906837</td><td></td></tr><tr><td>Sterile filtration unit, MILLEX-GV</td><td>Millipore</td><td>ML481051</td><td></td></tr><tr><td>SP8 AOBS confocal laser scanning microscope</td><td>Leica</td><td></td><td>Environmental chamber needed for temperature and CO&lt;sub&gt;2&lt;/sub&gt; control (Life Imaging Services)</td></tr><tr><td colspan="4">Key: IF, Immunofluorescence; WB, Western Blot</td></tr></tbody></table>

a rapid, scalable method for the isolation, functional study, and analysis of cell-derived extracellular matrix

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and pathological processes. However, the isolation of ECM, from tissues or cell culture, is complicated by the insoluble and cross-linked nature of the assembled ECM and by the potential contamination of ECM extracts with cell surface and intracellular proteins. Here, we describe a method for use with cultured cells that is rapid and reliably removes cells to isolate a cell-derived ECM for downstream experimentation.
Through use of this method, the isolated ECM and its components can be visualized by in situ immunofluorescence microscopy. The dynamics of specific ECM proteins can be tracked by tracing the deposition of a tagged protein using fluorescence microscopy, both before and after the removal of cells. Alternatively, the isolated ECM can be extracted for biochemical analysis, such as sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting. At larger scales, a full proteomics analysis of the isolated ECM by mass spectrometry can be conducted. By conducting ECM isolation under sterile conditions, sterile ECM layers can be obtained for functional or phenotypic studies with any cell of interest. The method can be applied to any adherent cell type, is relatively easy to perform, and can be linked to a wide repertoire of experimental designs.

The protocol described in steps 1.1-1.5 acts to remove cells and leave behind the cell-derived ECM. The protocol was carried out on COS-7 cells grown for 96 h on glass coverslips, and the cell-derived ECM was analyzed by fluorescence microscopy. The ammonium hydroxide treatment removed the cells effectively, as established by the loss of the cell nuclei and actin cytoskeleton, according to DAPI and phalloidin staining, respectively (Figure 1). The extraction of cells with 2 M urea was not as effective as ammonium hydroxide; traces of DAPI and phalloidin staining were detected after treatment. 8 M urea had a similar effect to ammonium hydroxide (Figure 1). Next, cell-derived ECM was visualized before and after the ammonium hydroxide treatment (steps 2.1-2.11). COS-7 cells were transfected with a monomeric red fluorescent protein (mRFP)-tagged thrombospondin-1 C-terminal trimer, (mRFPovTSP1C)11 and grown on a 35 mm dish with a gridded glass base.
Two hours post-plating, a suitable grid square that contained multiple healthy cells expressing mRFPovTSP1C was visualized under a confocal microscope. A reference phase contrast image was taken of this area, and then fluorescence time-lapse imaging was carried out every 1 min for 2 h (Figure 2A). Cells were then removed using ammonium hydroxide, and the same grid square on the dish was relocated under phase contrast and then re-analyzed by fluorescence microscopy. The cells were no longer present in the grid square, but mRFPovTSP1C remained within the ECM, detected by fluorescence microscopy as characteristic puncta (Figure 2A). Any mRFPovTSP1C deposited in the ECM prior to cell removal was identified by comparing the fluorescence pattern pre- and post-removal of cells. The fluorescent puncta identified in both images (Figure 2A, example arrowed) are inferred to be associated with the ECM.
Ammonium hydroxide treatment was then used to isolate native ECM from cultured, adherent cells. Human dermal fibroblasts (HDF) were grown on glass coverslips for 96 h. Cells were removed using ammonium hydroxide, and the ECM was probed with an anti-collagen I antibody, which demonstrated a characteristic fibrillar and meshwork staining pattern16 (Figure 2B). Fibronectin patterning was examined around fixed, non-permeabilized rat chondrosarcoma cells (RCS) and within the ECM after the removal of the RCS cells (Figure 2C). The ammonium hydroxide isolation of ECM was also carried out under sterile conditions so that &#34;test&#34; cells could be re-plated onto the ECM for phenotypic or functional studies. For example, &#34;test&#34; COS-7 cells were plated for 2 h onto sterile ECM produced by RCS cells, and then they were fixed, permeabilized, and analyzed for F-actin organization by FITC-phalloidin staining, in comparison to COS-7 cells producing their own ECM (steps 3.1-3.9) (Figure 3).
At a larger scale, the method was used to isolate ECM for biochemical procedures. For example, RCS cells were grown on two 100 mm cell culture dishes for 7 days, and the procedures in steps 4.1-4.7 were followed. Proteins in the cell-derived ECM were collected by scraping them into hot SDS-PAGE sample buffer and were separated by SDS-PAGE on a 7% polyacrylamide gel under reducing conditions. The gel was stained for protein, and the four major bands were each isolated and analyzed by mass spectrometry (Figure 4A). A number of ECM proteins, including fibronectin, thrombospondin 1 (TSP1), thrombospondin 5/cartilage oligomeric matrix protein (TSP5 / COMP), and matrilin-1 were identified (Figure 4B).
Alternatively, smaller-scale preparations of ECM can be evaluated by immunoblots. For example, cell-derived ECM from COS-7 cells ectopically expressing mRFPovTSP1C was separated by SDS-PAGE, transferred to a PVDF membrane, and probed for RFP with an anti-RFP antibody (Figure 4C). This technique is sensitive enough to detect differences in deposition between a native trimer of TSP1 (mRFPovTSP1C) and an engineered pentamer of the TSP1 C-terminal region (mRFP-TSP-5-1C)17 (Figure 4C). To investigate the endogenous ECM of HDF, the presence of specific proteins in cell lysate or the isolated ECM was examined by immunoblotting. The characteristic ECM proteins fibronectin and thrombospondin-1 were detected in the ECM, whereas the abundant intracellular proteins &#945;-tubulin and &#946;-actin were absent from the isolated ECM (Figure 4D).
<img alt="Figure 1" src="/files/ftp_upload/55051/55051fig1.jpg" /> 
Figure 1: Comparison of Methods of Cell Removal. COS-7 cells were grown at high density for 96 h on coverslips, fixed in 2% PFA, and permeabilized in 0.5% Triton X100, or were treated as indicated for cell removal. Coverslips were then stained with FITC-phalloidin to visualize F-actin and DAPI to visualize nuclei. Scale bar = 25 &#181;m. This figure is modified from an original publication in the Journal of Cell Science11. <a href="http://cloudflare.jove.com/files/ftp_upload/55051/55051fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55051/55051fig2.jpg" /> 
Figure 2: Identification of ECM Proteins in Isolated ECM. (A) Phase-contrast and fluorescence images of COS-7 cells expressing mRFPovTSP1C and grown on a 35 mm dish with a glass-bottomed, gridded base for 2 h. Images are shown before and after the removal of the cells by ammonium hydroxide. The edge of the grid letter is indicated in the phase-contrast images with a dotted line. White arrows indicate examples of fluorescent puncta that were present before and after cell removal. (B) Detection of collagen I in HDF ECM by indirect immunofluorescence. HDF were grown for 96 h and removed with ammonium hydroxide, and the ECM was stained for human fibrillar collagen I. The ECM was also stained with FITC-conjugated anti-rabbit secondary antibody alone. (C) Localization of fibronectin around RCS cells or within isolated RCS ECM, as detected by indirect immunofluorescence. RCS cells were grown for 48 h, fixed in 2% PFA, and stained for fibronectin and with DAPI. In parallel dishes, cells were removed with 20 mM ammonium hydroxide and the ECM was stained for fibronectin and with DAPI. Cells were also stained with FITC-conjugated anti-mouse IgG antibody alone. <a href="http://cloudflare.jove.com/files/ftp_upload/55051/55051fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55051/55051fig3.jpg" /> 
Figure 3: Use of Sterile ECM for Functional Studies. RCS &#34;matrix producer&#34; cells were grown for 48 h on glass coverslips and then treated with 20 mM ammonium hydroxide under sterile conditions to remove the cells. COS-7 &#34;test&#34; cells were plated on coverslips with or without isolated RCS ECM for 2 h, fixed in 2% PFA, permeabilized in 0.5% Triton X100, and stained with FITC-phalloidin to visualize F-actin and DAPI to visualize nuclei. COS-7 cells plated on RCS ECM spread more extensively and formed large microfilament bundles (example arrowed) and edge ruffles. <a href="http://cloudflare.jove.com/files/ftp_upload/55051/55051fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55051/55051fig4.jpg" /> 
Figure 4: Identification of Proteins from Isolated ECM by SDS-PAGE, Mass Spectrometry, or Immunoblotting. (A) RCS cells were grown for 7 days and removed with 20 mM ammonium hydroxide; the ECM was scraped up into hot SDS-PAGE sample buffer containing 100 mM DTT. The ECM preparation was separated by SDS-PAGE on a 7% polyacrylamide gel under reducing conditions. Four significant bands (arrows 1-4) were identified by protein staining. (B) Proteins from RCS ECM bands 1-4 were identified by MALDI mass spectrometry. (C) COS-7 cells expressing either mRFPovTSP1C (trimer) or mRFP-TSP-5-1C (pentamer) were cultured for 48 h and removed with 20 mM ammonium hydroxide, and the was ECM isolated into hot SDS-PAGE sample buffer containing 100 mM DTT. The ECM preparation was separated by SDS-PAGE on a 7% polyacrylamide gel under reducing conditions, transferred to a PVDF membrane, and probed with an anti-RFP antibody. This panel is modified from an original publication in Bioscience Reports17. (D) HDF were grown for 96 h and then treated either with 2% deoxycholic acid in PBS (cell lysate) or with 20 mM ammonium hydroxide to remove cells. The ECM was scraped into hot SDS-PAGE sample buffer. The two fractions were separated by SDS-PAGE on a 10% polyacrylamide gel under reducing conditions, transferred to a PVDF membrane, and probed with antibodies, as indicated. In each panel, the positions of molecular weight markers are indicated in kDa. <a href="http://cloudflare.jove.com/files/ftp_upload/55051/55051fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix at JoVE.com

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

The extracellular matrix plays a major role in defining the microenvironment of cells and in modulating cell behavior and phenotype. We describe a rapid method for the isolation of cell-derived extracellular matrix, which can be adapted to different scales for microscopic, biochemical, proteomic, or functional studies.

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and ...

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Universidad Científica del Sur

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16.5K Views.  University of Bristol. The extracellular matrix plays a major role in defining the microenvironment of cells and in modulating cell behavior and phenotype. We describe a rapid method for the isolation of cell-derived extracellular matrix, which can be adapted to different scales for microscopic, biochemical, proteomic, or functional studies. 

Video: A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

Analyzing the Communication Between Monocytes and Primary Breast Cancer Cells in an Extracellular Matrix Extract (ECME)-based Three-dimensional System

Embedded in the extracellular matrix (ECM), normal and neoplastic epithelial cells intimately communicate with hematopoietic and non-hematopoietic cells, thus greatly influencing normal tissue homeostasis and disease outcome. In breast cancer, tumor-associated macrophages (TAMs) play a critical role in disease progression, metastasis, and recurrence; therefore, understanding the mechanisms of monocyte chemoattraction to the tumor microenvironment and their interactions with tumor cells is important to control the disease. Here, we provide a detailed description of a three-dimensional (3D) co-culture system of human breast cancer (BrC) cells and human monocytes. BrC cells produced high basal levels of regulated on-activation, normal T-cell expressed and secreted (RANTES), monocyte chemoattractant protein-1 (MCP-1), and granulocyte-macrophage colony-stimulating factor (G-CSF), while in co-culture with monocytes, pro-inflammatory cytokines Interleukin (IL)-1 beta (IL-1&#946;) and IL-8 were enriched together with matrix metalloproteinases (MMP)-1, MMP-2, and MMP-10. This tumor stroma microenvironment promoted resistance to anoikis in MCF-10A 3D acini-like structures, chemoattraction of monocytes, and invasion of aggressive BrC cells. The protocols presented here provide an affordable alternative to study intra-tumor communication and are an example of the great potential that in vitro 3D cell systems provide to interrogate specific features of tumor biology related to tumor aggression.

Analyzing the Communication Between Monocytes and Primary Breast Cancer Cells in an Extracellular Matrix Extract ECME-based Three-dimensional System

Here, we describe a three-dimensional culture method to analyze the morphology of primary breast cancer cells, as well as to study their direct/indirect interactions with monocytes and the outcomes such as collagen degradation, immune cell recruitment, cell invasion, and promotion of cancer-related inflammation.

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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 ...

Bioengineering

A High-throughput Cell Microarray Platform for Correlative Analysis of Cell Differentiation and Traction Forces

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly controlled, high-throughput engineered system for measuring the impact of arrayed biochemical signals on cell differentiation. Recent efforts using cell microarrays have demonstrated their utility for combinatorial studies in which many microenvironmental factors are presented in parallel. However, these efforts have focused primarily on investigating the effects of biochemical cues on cell responses. Here, we present a cell microarray platform with tunable material properties for evaluating both cell differentiation by immunofluorescence and biomechanical cell&#8211;substrate interactions by traction force microscopy. To do so, we have developed two different formats utilizing polyacrylamide hydrogels of varying Young&#39;s modulus fabricated on either microscope slides or glass-bottom Petri dishes. We provide best practices and troubleshooting for the fabrication of microarrays on these hydrogel substrates, the subsequent cell culture on microarrays, and the acquisition of data. This platform is well-suited for use in investigations of biological processes for which both biochemical (e.g., extracellular matrix composition) and biophysical (e.g., substrate stiffness) cues may play significant, intersecting roles.

Cell differentiation is regulated by a host of microenvironmental factors, including both matrix composition and substrate material properties. We describe here a technique utilizing cell microarrays in conjunction with traction force microscopy to evaluate both cell differentiation and biomechanical cell&#8211;substrate interactions as a function of microenvironmental context.

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly ...

3D Models in Cancer Biology: Invasion Dynamics in Heterospheroids

Alternative Strategy to Analyze In Vitro Cell Invasion of 3D Cultures

Spheroid culture is a 3D model that provides an improved replication of the in vivo microenvironment compared to traditional two-dimensional (2D) cultures. Invasion is a cellular outcome of utmost interest in cancer biology. In this protocol, we have devised an alternative strategy for evaluating cancer cell invasion in vitro, employing heterospheroids comprised of oral squamous cell carcinoma (OSCC), cancer-associated fibroblasts (CAF), and monocytes. These heterospheroids aim to mimic the tumor microenvironment (TME), including two relevant non-neoplastic cell types alongside the cancer cells. Each cell type was labeled with vital fluorescent markers emitting in distinct wavelengths before spheroid formation. Once formed, heterospheroids were seeded onto a layer of human leiomyoma-derived extracellular matrix in the upper compartment of a microporous membrane. Invasion was assessed in the z-axis using confocal microscopy. Digital images were obtained in the corresponding fluorescent channels at 10 &#181;m intervals, covering a depth of 90 &#181;m in the z-axis. Analysis was performed using freeware image software by calculating the integrated fluorescence intensity in each image and fluorescence channel. This approach enables a more dynamic analysis of cell invasion patterns in a multilayered context, as well as the examination of spatial co-localization of different cell types during invasion.

Invasion is a major biological phenomenon in the development and progression of cancer. This process is influenced by non-neoplastic cells and components of the tumor microenvironment. The purpose of this study is to describe an alternative method for analyzing tumor cell invasion in vitro using three-dimensional (3D) cultures.

Spheroid culture is a 3D model that provides an improved replication of the in vivo microenvironment compared to traditional two-dimensional (2D) ...

Vasodilation of Isolated Vessels and the Isolation of the Extracellular Matrix of Tight-skin Mice

The interferon regulatory factor 5 (IRF5) is crucial for cells to determine if they respond in a pro-inflammatory or anti-inflammatory fashion. IRF5's ability to switch cells from one pathway to another is highly attractive as a therapeutic target. We designed a decoy peptide IRF5D with a molecular modeling software for designing small molecules and peptides.
IRF5D inhibited IRF5, reduced alterations in extracellular matrix, and improved endothelial vasodilation in the tight-skin mouse (Tsk/+). The Kd of IRF5D for recombinant IRF5 is 3.72 &plusmn; 0.74 x 10-6 M as determined by binding experiments using biolayer interferometry experiments. Endothelial cells (EC) proliferation and apoptosis were unchanged using increasing concentrations of IRF5D (0 to 100 &#181;g/mL, 24 h). Tsk/+ mice were treated with IRF5D (1 mg/kg/d subcutaneously, 21 d). IRF5 and ICAM expressions were decreased after IRF5D treatment. Endothelial function was improved as assessed by vasodilation of facialis arteries from Tsk/+ mice treated with IRF5D compared to Tsk/+ mice without IRF5D treatment. As a transcription factor, IRF5 traffics from the cytosol to the nucleus. Translocation was assessed by immunohistochemistry on cardiac myocytes cultured on the different cardiac extracellular matrices. IRF5D treatment of the Tsk/+ mouse resulted in a reduced number of IRF5 positive nuclei in comparison to the animals without IRF5D treatment (50 &#181;g/mL, 24 h). These findings demonstrate the important role that IRF5 plays in inflammation and fibrosis in Tsk/+ mice.

We describe the isolation of cardiac extracellular matrix from C57Bl/6J control mice, tight-skin mice, and tight-skin mice treated with the IRF5 inhibitory peptide. We also describe the vasodilation studies on the isolated vessels from C57Bl/6J, tight-skin mice and tight-skin mice treated with the IRF5 inhibitory peptide.

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Immunology and Infection

Fabrication of a Multiplexed Artificial Cellular MicroEnvironment Array

Cellular microenvironments consist of a variety of cues, such as growth factors, extracellular matrices, and intercellular interactions. These cues are well orchestrated and are crucial in regulating cell functions in a living system. Although a number of researchers have attempted to investigate the correlation between environmental factors and desired cellular functions, much remains unknown. This is largely due to the lack of a proper methodology to mimic such environmental cues in vitro, and simultaneously test different environmental cues on cells. Here, we report an integrated platform of microfluidic channels and a nanofiber array, followed by high-content single-cell analysis, to examine stem cell phenotypes altered by distinct environmental factors. To demonstrate the application of this platform, this study focuses on the phenotypes of self-renewing human pluripotent stem cells (hPSCs). Here, we present the preparation procedures for a nanofiber array and the microfluidic structure in the fabrication of a Multiplexed Artificial Cellular MicroEnvironment (MACME) array. Moreover, overall steps of the single-cell profiling, cell staining with multiple fluorescent markers, multiple fluorescence imaging, and statistical analyses, are described.

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3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and chemical complexity of living tissues and mimic in vivo tissue functions. Recent advances in microfabrication technologies have facilitated the development of 3D in vitro environments in which cells can be integrated into a well-defined extracellular matrix (ECM) and a defined set of soluble or matrix associated biomolecules. However, technological barriers have limited their widespread use in research laboratories. Here, we describe a method to construct simple devices for 3D culture and experimentation with cells and multicellular organoids in 3D microenvironments with a defined chemoattractant gradient. We illustrate the use of this platform for analysis of the response of epithelial cells and organoids to gradients of growth factors, such as epidermal growth factor (EGF). EGF gradients were stable in the devices for several days leading to directed branch formation in breast organoids. This analysis allowed us to conclude that collective gradient sensing by groups of cells is more sensitive vs. single cells. We also describe the fabrication method, which does not require photolithography facilities nor advanced soft lithography techniques. This method will be helpful to study 3D cellular behaviors in the context of the analysis of development and pathological states, including cancer.

We describe a method to construct devices for 3D culture and experimentation with cells and multicellular organoids. This device allows analysis of cellular responses to soluble signals in 3D microenvironments with defined chemoattractant gradients. Organoids are better than single cells at detection of weak noisy inputs.

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and ...

Isolation of Mammary Epithelial Cells from Three-dimensional Mixed-cell Spheroid Co-culture

While enormous efforts have gone into identifying signaling pathways and molecules involved in normal and malignant cell behaviors1-2, much of this work has been done using classical two-dimensional cell culture models, which allow for easy cell manipulation. It has become clear that intracellular signaling pathways are affected by extracellular forces, including dimensionality and cell surface tension3-4. Multiple approaches have been taken to develop three-dimensional models that more accurately represent biologic tissue architecture3. While these models incorporate multi-dimensionality and architectural stresses, study of the consequent effects on cells is less facile than in two-dimensional tissue culture due to the limitations of the models and the difficulty in extracting cells for subsequent analysis.
The important role of the microenvironment around tumors in tumorigenesis and tumor behavior is becoming increasingly recognized4. Tumor stroma is composed of multiple cell types and extracellular molecules. During tumor development there are bidirectional signals between tumor cells and stromal cells5. Although some factors participating in tumor-stroma co-evolution have been identified, there is still a need to develop simple techniques to systematically identify and study the full array of these signals6. Fibroblasts are the most abundant cell type in normal or tumor-associated stromal tissues, and contribute to deposition and maintenance of basement membrane and paracrine growth factors7.
Many groups have used three dimensional culture systems to study the role of fibroblasts on various cellular functions, including tumor response to therapies, recruitment of immune cells, signaling molecules, proliferation, apoptosis, angiogenesis, and invasion8-15. We have optimized a simple method for assessing the effects of mammary fibroblasts on mammary epithelial cells using a commercially available extracellular matrix model to create three-dimensional cultures of mixed cell populations (co-cultures)16-22. With continued co-culture the cells form spheroids with the fibroblasts clustering in the interior and the epithelial cells largely on the exterior of the spheroids and forming multi-cellular projections into the matrix. Manipulation of the fibroblasts that leads to altered epithelial cell invasiveness can be readily quantified by changes in numbers and length of epithelial projections23. Furthermore, we have devised a method for isolating epithelial cells out of three-dimensional co-culture that facilitates analysis of the effects of fibroblast exposure on epithelial behavior. We have found that the effects of co-culture persist for weeks after epithelial cell isolation, permitting ample time to perform multiple assays. This method is adaptable to cells of varying malignant potential and requires no specialized equipment. This technique allows for rapid evaluation of in vitro cell models under multiple conditions, and the corresponding results can be compared to in vivo animal tissue models as well as human tissue samples.

A simple method is described for analyzing effects of tissue fibroblasts on associated epithelial cells. The combination of this method and three-dimensional tissue culture can facilitate analysis of cells after isolation from 3D. The technique is applicable to cells of varying malignant potential, allowing systematic study of effects of tumor-associated stroma on tumor cells.

While enormous efforts have gone into identifying signaling pathways and molecules involved in normal and malignant cell behaviors1-2, much of this work ...

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.

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

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Isolation of Mammary Epithelial Cells from Three-dimensional Mixed-cell Spheroid Co-culture

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