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

The overall goal of this procedure is to isolate extracellular matrix from cultured cells for a wide variety of downstream experiments, to help understand functions and properties of extracellular matrix in normal tissues and disease processes. This method can help answer key questions in the extracellular matrix field, such as the organization, molecular composition, protein interactions or functional properties of extracellular matrix. Also the extracellular matrix could be prepared from any adherent cell type.

The main advantage of this technique is that they can be carried out in different scales to efficiently remove cells and retain extracellular matrix proteins. Demonstrating the procedure will be Silvia Rosini, a graduate student in my laboratory, and Andrew Hellewell a postdoctoral researcher. To begin, plate cells on cover slips for the imaging of live cells and extracellular matrix or ECM.

For florescence microscopy studies, a fixed ECM. Or for preparation of cell-derived ECM for small scale functional studies. Incubate the cells at an appropriate temperature and humidity for a suitable time for the cells to deposit ECM.

Typically greater than 16 hours. In an extractor hood, prepare 20 millimolar ammonium hydroxide in a suitable vessel by using deionized water to dilute the stock solution one to 14. Remove the cells from the incubator and gently remove the culture medium.

Then add PBS without calcium or magnesium by gently pouring against the wall of the dish. Rock the dish twice and pour away the liquid. Then repeat the wash two more times.

In the extractor hood, remove the last PBS wash, add 1.5 milliliters of ammonium hydroxide per 60 millimeter dish, and incubate the plates at room temperature for five minutes. Gently agitate the dishes every minute to ensure lysis of all the cells. Next, add copious amounts of deionized water to each dish and rock the plate.

Then pour away the ammonium hydroxide solubilized material, which is composed of ammonium hydroxide, lysed cells, and deionized water. Wash the insoluble ECM layer in copious amounts of deionized water four more times, to ensure the complete removal of all the ammonium hydroxide solubilized material. It is essential that the cells are fully removed with copious washes in deionized water.

If these steps are not performed adequately, cellular material may remain on the dish and contaminate downstream analysis. For examination of the ECM by immunofluorescence, remove the remaining deionized water, fix the ECM by adding 2%paraformaldehyde, abbreviated PFA, and incubate the dishes at room temperature for 10 minutes. Use PBS to wash each fix sample twice, and dispose of the solution in a liquid waste container before carrying out fluorescence microscopy according to the text protocol.

In a laminar flow hood prepare sterile solutions of PBS without calcium and magnesium, 20 millimolar ammonium hydroxide, and deionized water by passing each solution through a sterile 0.2 micrometer filter into an appropriate sterile container. While maintaining sterile technique, use sterile PBS to gently wash the matrix producer cells on cover slips three times. After removing the final PBS wash, add 1.5 milliliters of sterile ammonium hydroxide to the 60 millimeter dish and incubate at room temperature for five minutes with intermittent rocking.

Add copious amounts of sterile deionized water to the dish with rocking, pour away the cell lysate into an appropriate waste container. Repeat this wash four more times to ensure the complete removal of the solubilized material. After trypsinizing and pelleting the cells according to the text protocol, resuspend the test cells in fresh sterile medium, and use a hemocytometer to count the cells.

Then plate the appropriate number of test cells and culture medium onto the sterile isolated ECM on cover slips and incubate for two to three hours or for the time required. To examine the organization of the actin cytoskeleton fix the test cells in 2%PFA and stain them with FITC-Phalloidin to visualize the F-actin, and DAPI to visualize the nuclei. After culturing a large number of cells, then removing the cells and washing the ECM as demonstrated earlier in the video, tilt each plate at an angle to drain the residual water to the bottom of the dish and use a P200 pipette to carefully remove any remaining water until the plate is completely dry.

At the same time heat SDS-PAGE sample buffer containing 100 millimolar DDT to 95 degree celsius for two minutes, and add 200 microliters to each 100 millimeter plate to be examined by immunoblot. Use a cell scraper to gather the sample to one side of the dish, and use a 200 microliter pipette tip to transfer it into a tube. To analyze the ECM by mass spectrometry, prepare a more concentrated sample by collecting the SDS-PAGE sample buffer from the dish, reheating it to 95 degrees Celsius, and using the same aliquot across two to three additional plates.

Transfer any residual SDS-PAGE sample buffer extract from the cell scraper tip into the tube to minimize the loss of ECM material. For effective extraction of the isolated ECM into SDS-PAGE sample buffer it is essential that the sample buffer is heated to 95 degrees, contains a reducing agent, and that the plate is scraped extensively. Shown here are COS-7 cells grown on glass cover slips.

Ammonium hydroxide treatment removed the cells effectively as established by the loss of the cell nuclei and actin cytoskeleton. In this figure, COS-7 cells were transfected with RFP tag thrombospondin-1. And the ECM was visualized before and after ammonium hydroxide treatment.

The fluorescent puncta identified in both images are inferred to be associated with the ECM. In this experiment human dermal fibroblasts were removed with ammonium hydroxide and the ECM was probed with with anti-collagen 1 antibody, which demonstrated a characteristic fibular and meshwork staining pattern. As shown here, fibronectin staining was carried out on the ECM from ammonium hydroxide extracted rat chondrosarcoma cells.

In this example, test COS-7 cells were plated for two hours onto sterile ECM produced by RCS cells and analyzed for F-actin organization by FITC-Phalloidin staining. In comparison to COS-7 cells producing their own ECM. In this experiment ECM was isolated from rat chondrosarcoma cells and ECM derived proteins were then separated by SDS-PAGE.

And the four major bands were isolated and analyzed by mass spectrometry. Fibronectin, thrombospondin-1 and five, and matrilin-1 were identified. Finally, the endogenous ECM of human dermal fibroblasts was analyzed by immunoblotting.

Fibronectin and thrombospondin-1 were detected, whereas the intracellular proteins alpha-tubulin and beta-actin were absent. Once mastered, the removal of cells with ammonium hydroxide and extraction of ECM with hot SDS-PAGE sample buffer from three 100 millimeter dishes can be done in one hour if it is performed properly. While attempting this procedure it's important to remember to make fresh ammonium hydroxide working solutions each time for removing the cells.

It's also very important to use copious washes with deionized water to make sure to remove all the cells. Following this procedure are the methods such as proteomics analysis of extracted ECM can be performed in order to answer additional questions such as a relative abundance of ECM proteins produced by different cell types. After watching this video, you should have a good understanding of how to remove cells efficiently from cell culture dishes and how to extract ECM.

Don't forget that working with paraformaldehyde can be extremely hazardous and precautions such as working in an extractor hood and wearing personal protective equipment should always be taken while performing this procedure.