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 border="0" cellpadding="0" cellspacing="0" width="971">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:320px;">Antibody to COL1A1</td>
			<td style="width:123px;">Novus</td>
			<td style="width:107px;">NB600-408</td>
			<td style="width:423px;">Rabbit polyclonal. Reacts with other fibrillar collagens as well as collagen I.&#160; IF:&#160; 1/200&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Antibody to fibronectin</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">F3648</td>
			<td style="width:423px;">Rabbit polyclonal. WB: 1/600 for 1.5 hr. IF: 1/200</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Antibody to thrombospondin 1&#160;</td>
			<td style="width:123px;">ThermoFisher&#160;</td>
			<td style="width:107px;">MA5-13398</td>
			<td style="width:423px;">Mouse monoclonal clone A6.1.&#160; WB: 1/150 for 1.5 hr</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Antibody to RFP</td>
			<td style="width:123px;">Abcam</td>
			<td style="width:107px;">ab62341</td>
			<td style="width:423px;">Rabbit polyclonal. WB: 1/2000 for 2 hr</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Antibody to &#946;-actin</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">A1978</td>
			<td style="width:423px;">Mouse monoclonal clone AC-15. WB: 1/10000 for 1.5 hr&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Antibody to &#945;-tubulin</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">T9026</td>
			<td style="width:423px;">Mouse monoclonal clone DM1A. WB: 1/5000 for 1.5 hr</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Cell Tracker Green</td>
			<td style="width:123px;">ThermoFisher&#160;</td>
			<td style="width:107px;">C2925</td>
			<td style="width:423px;">Fluorescent cell marker. Use at 1 &#181;M for 30 min</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:320px;">Fluorescein isothiocyanate (FITC)-Phalloidin</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">P-5282</td>
			<td style="width:423px;">Stock = 50mg/ml in DMSO.1/50 for 45 min</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">FITC-conjugated goat anti-mouse IgG</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">F8771</td>
			<td style="width:423px;">IF: 1/50 for 1 hr</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">FITC-conjugated sheep anti-rabbit IgG</td>
			<td style="width:123px;">SIgma</td>
			<td style="width:107px;">F7512</td>
			<td style="width:423px;">IF: 1/50 for 1 hr</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:320px;">Horseradish peroxidase (HRP)-conjugated goat anti-mouse IgG</td>
			<td style="width:123px;">LI-COR</td>
			<td style="width:107px;">926-80010</td>
			<td style="width:423px;">WB: 1/50000 for 1 hr</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">HRP-conjugated goat anti-rabbit IgG</td>
			<td style="width:123px;">LI-COR</td>
			<td style="width:107px;">926-80011</td>
			<td style="width:423px;">WB: 1/100000 for 1 hr</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Vectorshield mounting medium with DAPI</td>
			<td style="width:123px;">Vector</td>
			<td style="width:107px;">H-1200</td>
			<td style="width:423px;">Store at 4 C in the dark</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Dulbecco&#39;s Modified Eagle&#39;s Medium</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">D6429</td>
			<td style="width:423px;">Warm before use</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:320px;">Fibroblast growth medium</td>
			<td style="width:123px;">PromoCell</td>
			<td style="width:107px;">C-23010</td>
			<td style="width:423px;">Warm before use, supplement with 50 &#181;g / mL ascorbic acid</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Phenol red-free DMEM</td>
			<td style="width:123px;">ThermoFisher&#160;</td>
			<td style="width:107px;">21063-029</td>
			<td style="width:423px;">Warm before use</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Trypsin-EDTA solution</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">T3924</td>
			<td style="width:423px;">Warm before use</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:320px;">Gridded glass-bottomed dish</td>
			<td style="width:123px;">MatTek</td>
			<td style="width:107px;">P35G-2-14-C-GRID</td>
			<td style="width:423px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:320px;">NH4OH, 28% - 30% &#160;solution</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">221228</td>
			<td style="width:423px;">Dilute to 20 mM in de-ionised water. Once opened store in the dark, tightly capped.&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Paraformaldehyde, 16 % solution</td>
			<td style="width:123px;">Alfa Aesar</td>
			<td style="width:107px;">43368</td>
			<td style="width:423px;">Dilute to 2 % (v/v) in PBS</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Deoxycholic acid</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">D2510</td>
			<td style="width:423px;">Use at 2 % (v/v) final concentration</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Triton X-100</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">T8787</td>
			<td style="width:423px;">Dilute to 0.5 % (v/v) final concentration</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">GelCode Blue stain reagent</td>
			<td style="width:123px;">ThermoFisher&#160;</td>
			<td style="width:107px;">24590</td>
			<td style="width:423px;">Protein stain for SDS-PAGE gels</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Precision Plus protein standards</td>
			<td style="width:123px;">Biorad</td>
			<td style="width:107px;">161-0374</td>
			<td style="width:423px;">Protein standards for SDS-PAGE gels</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Trans-Blot SD Semi Dry transfer cell</td>
			<td style="width:123px;">Biorad</td>
			<td style="width:107px;">1703940</td>
			<td style="width:423px;">Efficient transfer of high molecular weight proteins</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">PVDF transfer membrane</td>
			<td style="width:123px;">Millipore</td>
			<td style="width:107px;">ISEQ00010</td>
			<td style="width:423px;">Immobilon-PSQ&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Ponceau S stain</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:107px;">P7170</td>
			<td style="width:423px;">Reversible protein stain for PVDF membrane</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:320px;">WesternSure Enhanced chemiluminescence (ECL) substrate</td>
			<td style="width:123px;">LI-COR</td>
			<td style="width:107px;">926-80200</td>
			<td style="width:423px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">High performance chemiluminescence film</td>
			<td style="width:123px;">GE Healthcare</td>
			<td style="width:107px;">28906837</td>
			<td style="width:423px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;">Sterile filtration unit, MILLEX-GV</td>
			<td style="width:123px;">Millipore</td>
			<td style="width:107px;">ML481051</td>
			<td style="width:423px;"></td>
		</tr>
		<tr height="46">
			<td height="46" style="height:46px;width:320px;">SP8 AOBS confocal laser scanning microscope</td>
			<td style="width:123px;">Leica</td>
			<td style="width:107px;"></td>
			<td style="width:423px;">Environmental chamber needed for temperature and CO2 control (Life Imaging Services)</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:320px;"></td>
			<td style="width:123px;"></td>
			<td style="width:107px;"></td>
			<td style="width:423px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:320px;">Key: IF, Immunofluorescence; 
			WB, Western Blot</td>
			<td style="width:123px;"></td>
			<td style="width:107px;"></td>
			<td style="width:423px;"></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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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 ...

a-rapid-scalable-method-for-isolation-functional-study-analysis-cell

Nanyang Technological University

Research

JoVE Journal

Biology

16.4K 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

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

Badania wizualne Wrażliwość na prędkość i kierunek ruchu w Jaszczurki

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Avidity-based Extracellular Interaction Screening (AVEXIS) for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ensuring cohesion within multicellular organisms. Proteins predicted to form extracellular interactions are encoded by approximately a quarter of human genes1, but despite their importance and abundance, the majority of these proteins have no documented binding partner. Primarily, this is due to their biochemical intractability: membrane-embedded proteins are difficult to solubilise in their native conformation and contain structurally-important posttranslational modifications. Also, the interaction affinities between receptor proteins are often characterised by extremely low interaction strengths (half-lives &lt; 1 second) precluding their detection with many commonly-used high throughput methods2. 
Here, we describe an assay, AVEXIS (AVidity-based EXtracellular Interaction Screen) that overcomes these technical challenges enabling the detection of very weak protein interactions (t1/2 &le; 0.1 sec) with a low false positive rate3. The assay is usually implemented in a high throughput format to enable the systematic screening of many thousands of interactions in a convenient microtitre plate format (Fig. 1). It relies on the production of soluble recombinant protein libraries that contain the ectodomain fragments of cell surface receptors or secreted proteins within which to screen for interactions; therefore, this approach is suitable for type I, type II, GPI-linked cell surface receptors and secreted proteins but not for multipass membrane proteins such as ion channels or transporters.
The recombinant protein libraries are produced using a convenient and high-level mammalian expression system4, to ensure that important posttranslational modifications such as glycosylation and disulphide bonds are added. Expressed recombinant proteins are secreted into the medium and produced in two forms: a biotinylated bait which can be captured on a streptavidin-coated solid phase suitable for screening, and a pentamerised enzyme-tagged (&beta;-lactamase) prey. The bait and prey proteins are presented to each other in a binary fashion to detect direct interactions between them, similar to a conventional ELISA (Fig. 1). The pentamerisation of the proteins in the prey is achieved through a peptide sequence from the cartilage oligomeric matrix protein (COMP) and increases the local concentration of the ectodomains thereby providing significant avidity gains to enable even very transient interactions to be detected. By normalising the activities of both the bait and prey to predetermined levels prior to screening, we have shown that interactions having monomeric half-lives of 0.1 sec can be detected with low false positive rates3.

Avidity-based Extracellular Interaction Screening AVEXIS for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

AVEXIS is a high throughput protein interaction assay developed to systematically screen for novel extracellular receptor-ligand pairs involved in cellular recognition processes. It is specifically designed to detect transient protein interactions that are difficult to identify using other high throughput approaches.

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ...

A Rapid and Specific Microplate Assay for the Determination of Intra- and Extracellular Ascorbate in Cultured Cells

Vitamin C (ascorbate) plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. For instance, within the brain, ascorbate acts in a neuroprotective and neuromodulatory manner that involves ascorbate cycling between neurons and vicinal astrocytes - a relationship that appears to be crucial for brain ascorbate homeostasis. Additionally, emerging evidence strongly suggests that ascorbate has a greatly expanded role in regulating cellular and systemic iron metabolism than is classically recognized. The increasing recognition of the integral role of ascorbate in normal and deregulated cellular and organismal physiology demands a range of medium-throughput and high-sensitivity analytic techniques that can be executed without the need for highly expensive specialist equipment. Here we provide explicit instructions for a medium-throughput, specific and relatively inexpensive microplate assay for the determination of both intra- and extracellular ascorbate in cell culture.

Ascorbate plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. Here we describe a medium-throughput, specific and inexpensive microplate assay for the determination of both intra- and extracellular ascorbate in cell culture.

Vitamin C (ascorbate) plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. For instance, within ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of cell proliferation, survival, migration, etc. The ECM plays important roles in development and in diverse pathologies including cardio-vascular and musculo-skeletal diseases, fibrosis, and cancer. Thus, characterizing the composition of ECMs of normal and diseased tissues could lead to the identification of novel prognostic and diagnostic biomarkers and potential novel therapeutic targets. However, the very nature of ECM proteins (large in size, cross-linked and covalently bound, heavily glycosylated) has rendered biochemical analyses of ECMs challenging. To overcome this challenge, we developed a method to enrich ECMs from fresh or frozen tissues and tumors that takes advantage of the insolubility of ECM proteins. We describe here in detail the decellularization procedure that consists of sequential incubations in buffers of different pH and salt and detergent concentrations and that results in 1) the extraction of intracellular (cytosolic, nuclear, membrane and cytoskeletal) proteins and 2) the enrichment of ECM proteins. We then describe how to deglycosylate and digest ECM-enriched protein preparations into peptides for subsequent analysis by mass spectrometry.

This protocol describes a procedure for enriching ECM proteins from tissues or tumors and deglycosylating and digesting the ECM-enriched preparations into peptides to analyze their protein composition by mass spectrometry.

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of ...

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both mitochondrial (respiration) and non-mitochondrial (other redox) reactions consume oxygen, but these reactions can be easily distinguished by chemical inhibition of mitochondrial respiration. However, while mitochondrial oxygen consumption is an unambiguous and direct measurement of respiration rate2, the same is not true for extracellular acid production and its relationship to glycolytic rate 3-6. Extracellular acid produced by cells is derived from both lactate, produced by anaerobic glycolysis, and CO2, produced in the citric acid cycle during respiration. For glycolysis, the conversion of glucose to lactate- + H+ and the export of products into the assay medium is the source of glycolytic acidification. For respiration, the export of CO2, hydration to H2CO3 and dissociation to HCO3- + H+ is the source of respiratory acidification. The proportions of glycolytic and respiratory acidification depend on the experimental conditions, including cell type and substrate(s) provided, and can range from nearly 100% glycolytic acidification to nearly 100% respiratory acidification 6. Here, we demonstrate the data collection and calculation methods needed to determine respiratory and glycolytic contributions to total extracellular acidification by whole cells in culture using C2C12 myoblast cells as a model.

Glycolysis is a defining metabolic marker in multiple biological systems. Monitoring glycolysis by measuring the extracellular flux of H+ is common, but requires correction to be quantitative and unambiguous. Here, we demonstrate how to gather and correct extracellular flux data to distinguish between respiratory and glycolytic sources of extracellular acidification.

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both ...

Filtration Isolation of Nucleic Acids:  A Simple and Rapid DNA Extraction Method

FINA, filtration isolation of nucleic acids, is a novel extraction method which utilizes vertical filtration via a separation membrane and absorbent pad to extract cellular DNA from whole blood in less than 2 min. The blood specimen is treated with detergent, mixed briefly and applied by pipet to the separation membrane. The lysate wicks into the blotting pad due to capillary action, capturing the genomic DNA on the surface of the separation membrane. The extracted DNA is retained on the membrane during a simple wash step wherein PCR inhibitors are wicked into the absorbent blotting pad. The membrane containing the entrapped DNA is then added to the PCR reaction without further purification. This simple method does not require laboratory equipment and can be easily implemented with inexpensive laboratory supplies. Here we describe a protocol for highly sensitive detection and quantitation of HIV-1 proviral DNA from 100 &#181;l whole blood as a model for early infant diagnosis of HIV that could readily be adapted to other genetic targets.

We describe here a simple and rapid paper-based DNA extraction method of HIV proviral DNA from whole blood detected by quantitative PCR. This protocol can be extended for use in detecting other genetic markers or using alternative amplification methods.

FINA, filtration isolation of nucleic acids, is a novel extraction method which utilizes vertical filtration via a separation membrane and absorbent pad ...

Isolation of Intact, Whole Mouse Mammary Glands for Analysis of Extracellular Matrix Expression and Gland Morphology

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal architecture. Here, a small pocket below the skin was created using Mayo scissors, allowing separation of the glands within the subcutaneous tissue from the underlying peritoneum. Visualization of the glands was aided by the use of 3.5x-R surgical micro loupes. The pelt was inverted and pinned back allowing identification of the intact mammary fat pads. Each of the #4 abdominal glands was bluntly dissected by sliding the scalpel blade laterally between the subcutaneous layer and the glands. Immediately post-harvest, glands were placed in 10% neutral buffered formalin for subsequent tissue processing. Excision of the entire gland is advantageous because it primarily eliminates the risk of excluding important tissue-wide interactions between ductal epithelial cells and other microenvironmental cellular populations that could be missed in a partial biopsy. One drawback of the methodology is the use of serial sections from fixed tissues which limits analyses of ductal morphogenesis and protein expression to discrete locations within the gland. As such, changes in ductal architecture and protein expression in 3 dimensions (3D) is not readily obtainable. Overall, the technique is applicable to studies requiring whole intact murine mammary glands for downstream investigations such as developmental ductal morphogenesis or breast cancer.

Here, we present a protocol for the isolation of whole, intact mouse mammary glands to investigate extracellular matrix (ECM) expression and ductal morphology. Mouse #4 abdominal glands were extracted from 8-10 week old female nulliparous mice, fixed in neutral buffered formalin, sectioned and stained using immunohistochemistry for ECM proteins.

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal ...

Nanoparticle Tracking Analysis for the Quantification and Size Determination of Extracellular Vesicles

The physiological and pathophysiological roles of extracellular vesicles (EVs) have become increasingly recognized, making the EV field a quickly evolving area of research. There are many different methods for EV isolation, each with distinct advantages and disadvantages that affect the downstream yield and purity of EVs. Thus, characterizing the EV prep isolated from a given source by a chosen method is important for interpretation of downstream results and comparison of results across laboratories. Various methods exist for determining the size and quantity of EVs, which can be altered by disease states or in response to external conditions. Nanoparticle tracking analysis (NTA) is one of the prominent technologies used for high-throughput analysis of individual EVs. Here, we present a detailed protocol for quantification and size determination of EVs isolated from mouse perigonadal adipose tissue and human plasma using a breakthrough technology for NTA representing major advances in the field. The results demonstrate that this method can deliver reproducible and valid total particle concentration and size distribution data for EVs isolated from different sources using different methods, as confirmed by transmission electron microscopy. The adaptation of this instrument for NTA will address the need for standardization in NTA methods to increase rigor and reproducibility in EV research.

We demonstrate how to use a novel nanoparticle tracking analysis instrument to estimate the size distribution and total particle concentration of extracellular vesicles isolated from mouse perigonadal adipose tissue and human plasma.

The physiological and pathophysiological roles of extracellular vesicles (EVs) have become increasingly recognized, making the EV field a quickly evolving ...