J.D.M. was supported by a PhD fellowship from INSERM/R&#233;gion Aquitaine and is now supported by a post-doctoral ARC fellowship and the Tisch Cancer Institute at the Mount Sinai School of Medicine. E.H. is supported by a PhD from the Minist&#232;re de l'Enseignement Sup&#233;rieur et de la Recherche. Z.E. is supported by a post-doctoral fellowship from Agence Nationale de la Recherche (ANR). C.M. is supported by the Tisch Cancer Institute at the Mount Sinai School of Medicine and J.J.B.C. is supported by the NIH/NCI grant K22CA196750, the TCI Young Scientist Cancer Research Award JJR Fund, and the Tisch Cancer Institute at Mount Sinai School of Medicine. This work was supported by a grant from ANR-13-JJC-JSV1-0005. F.S. is supported by the "Ligue nationale contre le cancer." V.M. and F.S. are supported by funding from "Equipe Labellis&#233;e Ligue Nationale contre le Cancer 2016" and Institut National du Cancer, INCA_8036, and PLBio2012.

NOTE: Prior to fixation, all steps are completed in a sterile laminar flow hood.
1. 2D matrix
NOTE: 2D matrices are used to determine the percentage of cells forming invadosomes and the matrix degradation area per cell.
<img alt="Figure 1" src="/files/ftp_upload/54911/54911fig1.jpg" /> 
Figure 1: Protocol. Summary of the protocol used to prepare gelatin and collagen I matrices. It is possible to make the gelatin matrix only or a gelatin and collagen I mixed matrix. <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Gelatin matrix 
	NOTE: Fluorescent gelatin matrices are used to quantify matrix degradation, and non-fluorescent gelatin is used to promote the adhesion of collagen I fibers. Fluorescent gelatin matrices are also used for podosome and invadopodia studies (see Figure 1).
	<ol>
		<li>Under a sterile laminar flow hood, place a piece of parafilm (20 cm x 15 cm) on the work area and disinfect it with 70% ethanol.</li>
		<li>Aliquot 20 &#181;L droplets of 1 mg/mL gelatin in a line, spaced approximately 1 cm apart; refer to Figure 1 for droplet organization.</li>
		<li>Cover each 20 &#181;L gelatin droplet with an autoclaved glass coverslip (12 mm in diameter).</li>
		<li>Incubate for 20 min at room temperature. If the gelatin is fluorescent, protect it from light.</li>
		<li>Aliquot 40 &#181;L droplets of 0.5% glutaraldehyde in 1x phosphate-buffered saline (PBS) in a line, each spaced approximately 1 cm below the line of coverslips from step 1.1.3. 
		CAUTION: Glutaraldehyde is a toxic fixative agent. Use gloves and do not inhale.</li>
		<li>Using jeweled forceps, transfer each gelatin-coated coverslip to a glutaraldehyde droplet. 
		NOTE: It is normal that a small gelatin droplet remains on the parafilm. Do not reuse this for another experiment.</li>
		<li>Incubate for 40 min at room temperature. For fluorescent coverslips, protect them from light.</li>
		<li>Take a 24-well cell culture plate and fill each well with 500 &#181;L of sterile 1x PBS. Place the coverslips, gelatin side up, into each well.</li>
		<li>Wash twice in 1x PBS for 5 min each. 
		NOTE: At this step, the coverslips can be used for experimentation or stored in 1x PBS at 4 &#176;C. Non-fluorescent coverslips can be stored for one week, and fluorescent coverslips can be stored for 48 h.</li>
	</ol>
	</li>
	<li>Fibrillar collagen type I matrix 
	NOTE: Fibrillar collagen type I matrix is used to study linear invadosome formation and the associated matrix degradation. A combination of fluorescent collagen I and non-fluorescent gelatin coverslips is used to colocalize invadosome markers with collagen fibers (e.g., F-actin in the far-red channel, collagen I in the red channel, Tks5 in the green channel, and Hoechst dye for a nuclear stain). On the other hand, a combination of non-fluorescent collagen I with non-fluorescent gelatin coverslips is used to maximize the number of invadosome markers stained (e.g., F-actin in the far-red channel, cortactin in the red channel, Tks5 in the green channel, and Hoechst dye for a nuclear stain). Finally, a combination of non-fluorescent collagen I with fluorescent gelatin coverslips is used to quantify the gelatin-associated degradation (e.g., F-actin in the far-red channel, Tks5 in the red channel, in the green channel, and Hoechst dye for a nuclear stain) (see Figure 1). Hoechst dye is used to stain the nucleus, and it also checks for mycoplasma contamination, in addition to a PCR test done regularly on the cell lines used.
	<ol>
		<li>Fluorescent collagen I
		<ol>
			<li>Use commercial collagen I extracted from rat-tail tendon in 0.02 N acetic acid. Withdraw the quantity of collagen I necessary to prepare 500 &#181;L of solution per coverslip at a final concentration of 0.4 mg/mL of collagen I.</li>
			<li>Add 5-carboxy x rhodamine succinimidyl ester at a final concentration of 10 &#181;g/mL, calculated for the final volume of collagen I solution (n x 500 &#181;L).</li>
			<li>Mix the solution by pipetting up and down and incubate for 5 min at room temperature, protected from light.</li>
			<li>Dilute to the final volume with 1X Dulbecco&#39;s Phosphate-Buffered Saline (DPBS) containing calcium and magnesium, and use directly.</li>
		</ol>
		</li>
		<li>Fibrillar collagen I coverslip
		<ol>
			<li>Use the non-fluorescent or fluorescent gelatin coverslips previously prepared in step 1.1.</li>
			<li>Prepare a solution of fluorescent or non-fluorescent collagen I at a final concentration of 0.4 mg/mL in 1X DPBS containing calcium and magnesium.</li>
			<li>Add 500 &#181;L of fluorescent or non-fluorescent collagen I solution per coverslip in a 24-well cell culture plate.</li>
			<li>Incubate for 4 h at 37 &#176;C for collagen I polymerization.</li>
			<li>After incubation, remove the excess collagen I by tilting the plate and pipetting on the side of the well (not directly on the coverslip) to avoid damaging the matrix. 
			NOTE: This matrix cannot be stored. Seed the cells immediately after removing excess collagen I.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Cell seeding, time of culture, and fixation
<ol>
	<li>Depending on the cell type, prepare the cells in the appropriate culture medium and add cells for a final volume of 500 &#181;L per well.</li>
	<li>To characterize the ability of a cell line to form linear invadosomes and to degrade the matrix, plate the cells for 4 h, 12 h, and 24 h on collagen I matrix to determine the kinetics of linear invadosome formation and the associated degradation activity. 
	NOTE: Table 1 contains examples of cell lines tested in the laboratory.
	<table border="1">
		<tbody>
			<tr>
				<td>Cell line</td>
				<td>Number of cells per coverslip</td>
				<td>Time of contact with matrix for&#160; linear invadosome quantification</td>
				<td>Time of contact with matrix for&#160; quantification of&#160; linear invadosome degradation activity</td>
			</tr>
			<tr>
				<td>MDA MB 231</td>
				<td>40,000</td>
				<td>4 h</td>
				<td>12 h</td>
			</tr>
			<tr>
				<td>HUH7</td>
				<td>40,000</td>
				<td>12 h</td>
				<td>24 h&#160;</td>
			</tr>
			<tr>
				<td>HEP 3B</td>
				<td>40,000</td>
				<td>12 h</td>
				<td>24 h&#160;</td>
			</tr>
			<tr>
				<td>SNU 398</td>
				<td>40,000</td>
				<td>12 h</td>
				<td>24 h&#160;</td>
			</tr>
			<tr>
				<td>NIH 3T3 src</td>
				<td>30,000</td>
				<td>4 h</td>
				<td>12 h</td>
			</tr>
			<tr>
				<td>A431</td>
				<td>50,000</td>
				<td>6 h</td>
				<td>24 h&#160;</td>
			</tr>
			<tr>
				<td>A549</td>
				<td>40,000</td>
				<td>12 h</td>
				<td>24 h&#160;</td>
			</tr>
			<tr>
				<td>RAW</td>
				<td>40,000</td>
				<td>12 h</td>
				<td>12 h</td>
			</tr>
			<tr>
				<td>PAE</td>
				<td>40,000</td>
				<td>12 h</td>
				<td>12 h</td>
			</tr>
		</tbody>
	</table>
	Table 1: Cell seeding and incubation time reference. This table presents a non-exhaustive list of cell lines used to study linear invadosomes. It indicates the number of cells to seed on the matrix, the time necessary to observe linear invadosomes, and the time necessary to observe the associated matrix degradation. Linear invadosomes have a short lifetime. To be able to observe linear invadosomes in the maximum number of cells, the cells have to be in contact with the collagen I matrix for a short period of time. On the other hand, to be able to visualize the reported gelatin degradation, the cells must be in contact with the collagen I matrix for a longer period of time than those listed above.</li>
	<li>After the removal of the cell culture medium, fix the coverslips for 10 min in 500 &#181;L of 4% paraformaldehyde in 1X PBS.</li>
	<li>Wash twice with 1x PBS.</li>
	<li>Store at 4 &#176;C, protected from light.</li>
</ol>
3. Immunofluorescence
<ol>
	<li>Permeabilize the cells for 10 min with a freshly prepared solution of 0.2% Triton X-100 in 1x PBS. Do not reuse this solution.</li>
	<li>Wash the coverslips twice with 1x PBS.</li>
	<li>Place a piece of parafilm (20 cm x 15 cm) on the bench.</li>
	<li>Dilute primary antibody in 4% bovine serum albumin (BSA) in 1x PBS, as described in the Materials Table.</li>
	<li>Aliquot 50 &#181;L droplets of primary antibody solution in a line, each spaced approximately 1 cm apart.</li>
	<li>Place each coverslip on a droplet.</li>
	<li>Incubate for 40 min at room temperature, protected from light.</li>
	<li>After the incubation period, wash twice with 1x PBS.</li>
	<li>Dilute the secondary antibody, phalloidin, and Hoechst dye in 4% BSA in 1x PBS.</li>
	<li>Aliquot 50 &#181;L droplets of the mix, each spaced approximately 1 cm below the first line of coverslips, as done previously.</li>
	<li>Place each coverslip on a droplet.</li>
	<li>Incubate for 30 min at room temperature, protected from light.</li>
	<li>After incubation, wash twice with 1x PBS.</li>
	<li>Wash once with distilled water to remove the salts.</li>
	<li>Mount the coverslips on a glass slide with polymerizing mounting medium.</li>
	<li>Let the coverslips dry overnight at room temperature, protected from light, before microscopy.</li>
</ol>
4. Quantifications
<img alt="Figure 2" src="/files/ftp_upload/54911/54911fig2.jpg" /> 
Figure 2: Linear invadosome quantification. The macro requires confocal images of F-actin and Tks5 staining (format: 1,024 x 1,024). (A) First, open the F-actin picture and make a mask. (B) Then, open the Tks5 picture and threshold it. Apply the macro. (C) Analyzed results will appear in two tables. The number of linear invadosomes per cell and other information, such as the percentage area used by linear invadosomes in the cell, will be in the Summary table. The size of each linear invadosome will be in the Results table. <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Percentage of cells forming linear invadosomes
	<ol>
		<li>Count the cells forming linear invadosomes, approximately 100 cells per coverslip (use technical triplicates). Quantify at least 300 cells per n, with n = 3.</li>
		<li>Calculate the percentage of cells forming linear invadosomes by taking the average of three independent experiments, resulting in 900 quantified cells.</li>
	</ol>
	</li>
	<li>Linear invadosome size and number per cell 
	NOTE: Use a confocal microscope to take sequential images of the F-actin and Tks5 stains. The macro is adapted to a 1,024 x 1,024 image format.
	<ol>
		<li>Use ImageJ with the supplementary macro.</li>
	</ol>
	</li>
	<li>Degradation per cell
	<ol>
		<li>Use three coverslips per condition.</li>
		<li>Take 30 images per coverslip of fluorescent gelatin and Hoechst-stained nuclei.</li>
		<li>Use the ImageJ software as described in Martin KH et al.9.</li>
	</ol>
	</li>
</ol>
5. 3D Collagen Invasion Assay
<img alt="Figure 3" src="/files/ftp_upload/54911/54911fig3.jpg" /> 
Figure 3: Invasion assay schema. Summary of the invasion assay protocol. Collagen I is cross-linked with succinimidyl ester dye and polymerized for 1 h at 37 &#176;C. Cells are added in serum-free medium on top of the collagen I plug. The insert is placed in medium with 10% FBS. Collagen I plugs are fixed 1 h after cell seeding in the control and 3 days after cell seeding in the experimental condition to determine the invasion capacity of the cells. <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Collagen I plug preparation
	<ol>
		<li>Make a time course of 4 h, 12 h, 24 h, 48 h, and 72 h for the first experiment to determine the kinetics of invasion into the gel according to the cell type. For each experiment, add a 1 h time point as a reference time point when no invasion occurs.</li>
		<li>Prepare the collagen I solution on ice under a sterile laminar flow hood. Prepare 500 &#181;L of solution for 3 Boyden chamber inserts.</li>
		<li>Aspirate 1 mg of commercial rat-tail collagen I to prepare a solution with a final concentration of 2 mg/mL collagen I.</li>
		<li>Add 5-carboxy x Rhodamine succinimidyl ester at a final concentration of 1 &#181;g/mL, calculated for the final volume of collagen I solution (in this case, 500 &#181;L).</li>
		<li>Mix well and incubate for 5 min on ice under the sterile laminar flow hood, protected from light.</li>
		<li>Add 50 &#181;L of ice-cold, filtered 10X PBS and 6.25 &#181;L of ice-cold 1 M sodium hydroxide (NaOH).</li>
		<li>Add sterile water according to the formula 443.75 - x &#181;L of collagen I.</li>
		<li>Add 100 &#181;L of solution per Boyden chamber insert.</li>
		<li>Incubate for 1 h at 37 &#176;C to allow for polymerization.</li>
		<li>Add cell culture medium enriched with fetal bovine serum (FBS) into new wells; place inserts into these wells.</li>
		<li>Seed 30,000 cells on top of the collagen I plug in FBS-free medium.</li>
		<li>After culturing the cells at 37 &#176;C, fix the collagen I plug for 30 min in 4% paraformaldehyde in 1x PBS.</li>
		<li>Complete immunofluorescence as in step 3, with the exception of the following steps: permeabilize with 0.2% Triton X-100 in 1x PBS for 30 min, not 10 min; incubate with primary and secondary antibody solutions for 1.5 h each, rather than 40 min and 30 min, respectively. Stain for F-actin or nuclei to localize the cells.</li>
		<li>After immunofluorescence, use a scalpel to cut around the membrane of the insert and carefully transfer the collagen I plug to a glass-bottom dish. Make sure that the top of the collagen I plug is in contact with the glass. Keep the insert membrane attached to the bottom of the collagen I plug in order to maintain the integrity of the plug. 
		NOTE: The collagen I plug remains intact in the glass-bottom dish. The depth between the glass bottom and the plastic surrounding the glass bottom is equivalent to the thickness of the plug.</li>
		<li>Place a coverslip on the collagen I plug and mount with polymerizing mounting medium to prevent the plug from drying out.</li>
		<li>Use an inverted confocal microscope to image the collagen I plug; the top of the plug is in contact with the glass bottom of the dish.
		<ol>
			<li>Acquire a z-stack from the top to the bottom of the collagen I plug to visualize cell invasion. Determine the thickness of the z-stack by examining how deeply the cells invade the imaging field. Perform acquisitions with a 40X oil objective in a 512 x 512 format with a z-step size of 0.25 &#956;m. 
			NOTE: For reference, the average z-stack for MDA-MB-231 cells 3 days after seeding is 30 &#181;m.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Classically, invadosomes are studied in vitro without regard to the microenvironment and the matrix on which the cells are plated. Several types of matrices are currently used, including gelatin, fibronectin, vitronectin, or high-density fibrillar collagen (HDFC)7,11; however, these are often not representative of the microenvironment in which cells reside and are not physiologically relevant. Here, a novel type of matrix, which consists of an association between gelatin and fibrillar type I collagen, was used. The use of type I collagen fibrils allows us to highlight a novel class of invadosomes, known as linear invadosomes, which specifically form on collagen I in its physiological architecture7. Labeling the collagen I allows us to observe linear invadosomes along the fibers and to quantify their formation using cellular markers such as Tks5 and cortactin. Using the mixed matrix, we have identified a new receptor, the discoidin domain receptor 1 (DDR1), involved in the formation of linear invadosomes8.
The 2D mixed matrix allows us to quantify the matrix degradation activity of linear invadosomes via the zymography in situ assay. This assay reports the proteolysis activity of MMPs by analyzing the presence of black holes in the fluorescent gelatin layer. Interestingly, the organization of F-actin into linear invadosomes increases the matrix degradation activity compared to gelatin-only7. One limitation of this assay is that the gelatin is a non-physiological matrix, and it has been demonstrated that a more physiological matrix changes the cellular response to the microenvironment. An additional limitation exists in the way in which MMP activity is quantified on the gelatin-collagen I mixed matrices. Only the gelatin degradation that localizes under the collagen I fibril layer can be quantified; therefore, this is an indirect method to quantify collagen I matrix degradation. An alternative method to visualize the degradation activity of linear invadosomes is to label the collagen I fibers with a specific antibody against collagen cleavage sites (Col1-3/4C, immunoglobulin). This allows for the visualization of cleaved fibers by immunofluorescence. However, due to cell migration, this antibody is not very specific in 2D for degradation due to linear invadosome formation. Another way to visualize and quantify the degradation activity of the collagen I fibers is to use multiphoton microscopy and second harmonic generation8. This method allows for the imaging of collagen I fibers without any staining.
Our method of polymerizing type I collagen is different compared to other methods used in the literature12,13. For example, Artym et al. centrifuge coverslips coated with an alpha helix collagen I solution and allow for a brief polymerization of 30 min. This high-density fibrillar collagen (HDFC) is much stiffer and denser than our collagen I fibers but is less polymerized. This difference in matrix structure shifts the invadosome shape from linear to dot-like. The time, pH, concentration, and temperature are parameters that have to be controlled for ECM polymerization. For example, the lower the temperature of polymerization, the larger the fibers will be.
To study the involvement of linear invadosomes in cell invasion, the 3D collagen I plug assay was used. The 3D collagen I plug is already used by the scientific community in order to study invasive structures or the involvement of metalloproteinases in the invasion process14,15,16. These types of 3D matrices have limits regarding their rigidity-for example, the 3D collagen I plug is less rigid than the 2D matrix. Moreover, the type and origin of the collagen I is also important regarding the different organization of collagen fibers in vivo17. Finally, in regards to the microenvironment composition in vivo, only one element of the extracellular matrix, the type I collagen, was focused on. Further study is necessary to determine the relevance of linear invadosomes in vivo.
In addition to the study of linear invadosomes, the mixed matrix, described herein, could be used with other types of matrix components, such as fibronectin, vitronectin, and other types of collagens (e.g., type IV collagen). This protocol could be adapted, depending on the types of matrix elements that are of interest and the processes to be studied. However, it is clear that generating complex and physiological matrices will allow for the identification of new pathways involved in cell adhesion, migration, and invasion.

Extracellular matrix (ECM) remodeling occurs during physiological and pathological processes, such as angiogenesis and tumor cell invasion. In physiological conditions, many cell types, mostly from the hematopoietic lineage, are able to degrade ECM elements. For example, macrophages are able to cross anatomical barriers to reach tissues, and osteoclasts degrade bone matrix to ensure calcium homeostasis. More globally, all matrices in the body renew to maintain their physical and chemical properties. In cancer tissues, the tumor microenvironment composition is altered. For example, in breast and lung cancer, type I collagen is overexpressed and accumulates around the tumor. Moreover, this accumulation is associated with an increased risk of developing metastasis1,2. Cancer metastasis is dependent on the ability of cancer cells to degrade the ECM and invade adjacent tissues.
The invasive activity of cells is attributed to specialized actin-rich structures known as invadosomes. This term includes podosomes and invadopodia, which are present, respectively, in normal and cancer cells (e.g., macrophages, endothelial cells, and cancer cells such as the MDA-MB-231 breast cancer cell line). In vitro, invadosomes can organize into different shapes: dots, aggregates, or rosettes3. Classically, invadosomes are composed of an F-actin core containing several proteins, such as the scaffold protein Tks5 and cortactin, surrounded by adhesion plaque molecules, such as integrins and vinculin4. Recently, it was shown that Tks5 and the RhoGTPase Cdc42 can be used as a minimum molecular signature for functional invadosomes5. These structures are able to degrade the ECM via the recruitment and activation of specific metalloprotease proteins, such as MT1-MMP and MMP-26. In a previous study, we reported that the interaction between type I collagen fibrils and the discoidin domain receptor 1 (DDR1), a specific receptor of type I collagen fibrils, leads to the formation of a new class of invadosomes, named linear invadosomes. Linear invadosomes are formed along type I collagen fibrils7. These structures are able to degrade type I collagen fibrils via the recruitment and activation of MT1-MMP/MMP2. Moreover, their formation is dependent on Tks5 and Cdc425. In vitro, we demonstrated the existence of linear invadosomes and the involvement of DDR1 in their formation and activity8 using different strategies consisting of a combination of various matrix elements, including: i) fluorescent gelatin-coated coverslips, ii) fluorescent type I collagen fibril-coated coverslips, and iii) 3D type I collagen plugs. Due to the use of different matrices, we were able to study and characterize linear invadosome formation and their degradation activity7,8.
Finally, to better understand the biology of invasive cells, a combination of ECM components in both 2D and 3D culture systems were used, mimicking the complexity of the tumor microenvironment composition. Below are protocols for coating coverslips with gelatin/type I collagen in 2D and 3D culture systems.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Gelatin solution</td>
			<td>Sigma</td>
			<td>G1393</td>
			<td>Stock 20 mg/ml used at 1 mg/ml</td>
		</tr>
		<tr>
			<td>Gelatin from pig skin Oregon green 488 conjugate</td>
			<td>Molecular probe life technologies</td>
			<td>G13186</td>
			<td>5mg dilute at 1 mg/ml in sterile water</td>
		</tr>
		<tr>
			<td>Microscope cover glasses</td>
			<td>Marlenfeld GmbH &#38; Co</td>
			<td>111520</td>
			<td>&#216;12mm No.1</td>
		</tr>
		<tr>
			<td>2.5% Glutaraldehyd in 0.1M sodium cacodylate buffer pH 7.4</td>
			<td>Electron microscopy science</td>
			<td>15960</td>
			<td>Dilute at 0.5 % in sterile water</td>
		</tr>
		<tr>
			<td>1X PBS pH 7.4</td>
			<td>Gibco by life technologies</td>
			<td>10010-015</td>
			<td>Use this PBS in all steps before fixation</td>
		</tr>
		<tr>
			<td>Collagen I Rat tail</td>
			<td>Corning</td>
			<td>354236</td>
			<td></td>
		</tr>
		<tr>
			<td>5 carboxy X rhodamin siccinimidyl ester</td>
			<td>Life technologies</td>
			<td>C-6125</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS 1X + calcium + magnesium</td>
			<td>Gibco by life technologies</td>
			<td>14040-091</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde 16% solution</td>
			<td>Electron microscopy science</td>
			<td>15710</td>
			<td>Dilute at 4% in 1X PBS</td>
		</tr>
		<tr>
			<td>Triton X 100</td>
			<td>Sigma</td>
			<td>T9284</td>
			<td></td>
		</tr>
		<tr>
			<td>10X PBS buffer pH 7.4</td>
			<td>Ambion</td>
			<td>AM9625</td>
			<td>Dilute at 1X in water Use in steps after fixation</td>
		</tr>
		<tr>
			<td>Tks5 antibody</td>
			<td>Santa Cruz</td>
			<td>sc-30122</td>
			<td>Invadosome markers Dilution : 1/100</td>
		</tr>
		<tr>
			<td>Cortactin 4F11 antibody</td>
			<td>Millipore</td>
			<td>5180</td>
			<td>Invadosome markers Dilution :1/100</td>
		</tr>
		<tr>
			<td rowspan="2">DDR1 antibody</td>
			<td rowspan="2">Cell signaling</td>
			<td rowspan="2">5583</td>
			<td>Linear invadosome receptor</td>
		</tr>
		<tr>
			<td>Dilution :1/100</td>
		</tr>
		<tr>
			<td>Phalloidin FluoProbes</td>
			<td>Interchim</td>
			<td>FT-AZ0330</td>
			<td>Fibrillary actin marker Dilution :1/200</td>
		</tr>
		<tr>
			<td>Hoechst</td>
			<td>Sigma</td>
			<td>33258</td>
			<td>Nucleus marker Dilution :1/200</td>
		</tr>
		<tr>
			<td>Secondary antibodies FluoProbes</td>
			<td>Interchim</td>
			<td>FP-488 FP-547H or FP-647H</td>
			<td></td>
		</tr>
		<tr>
			<td>Albumin from bovine serum</td>
			<td>Sigma</td>
			<td>A2153</td>
			<td>Dilute at 4% in 1X PBS</td>
		</tr>
		<tr>
			<td>Fluoromount G mounting medium</td>
			<td>Interchim</td>
			<td>FP 483331</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ software</td>
			<td>Public domain</td>
			<td></td>
			<td>http://www.macbiophotonics.ca/imagej/</td>
		</tr>
		<tr>
			<td>Cell culture insert</td>
			<td>Corning</td>
			<td>353097</td>
			<td>8.0&#181;m pore size / 24 wells</td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sigma</td>
			<td>221465</td>
			<td></td>
		</tr>
	</tbody>
</table>

2d and 3d matrices to study linear invadosome formation and activity

Cell adhesion, migration, and invasion are involved in many physiological and pathological processes. For example, during metastasis formation, tumor cells have to cross anatomical barriers to invade and migrate through the surrounding tissue in order to reach blood or lymphatic vessels. This requires the interaction between cells and the extracellular matrix (ECM). At the cellular level, many cells, including the majority of cancer cells, are able to form invadosomes, which are F-actin-based structures capable of degrading ECM. Invadosomes are protrusive actin structures that recruit and activate matrix metalloproteinases (MMPs). The molecular composition, density, organization, and stiffness of the ECM are crucial in regulating invadosome formation and activation. In vitro, a gelatin assay is the standard assay used to observe and quantify invadosome degradation activity. However, gelatin, which is denatured collagen I, is not a physiological matrix element. A novel assay using type I collagen fibrils was developed and used to demonstrate that this physiological matrix is a potent inducer of invadosomes. Invadosomes that form along the collagen fibrils are known as linear invadosomes due to their linear organization on the fibers. Moreover, molecular analysis of linear invadosomes showed that the discoidin domain receptor 1 (DDR1) is the receptor involved in their formation. These data clearly demonstrate the importance of using a physiologically relevant matrix in order to understand the complex interactions between cells and the ECM.

Using a combination of two types of matrices, gelatin and type I collagen (Figures 1 and 4), we have highlighted a new type of invadosome, known as linear invadosomes. The labelling of these matrices allows for the observation of linear invadosome formation along collagen I fibers and of their degradation capabilities (Figure 5). The number of invasive structures can then be quantified by the macro described previously (step 4.2), and the degradation activity can also be determined using the ImageJ software, as describe by Martin et al. and Diaz et al.9,10. The mixed matrix allows for the characterization of linear invadosomes by defining DDR1 as the receptor necessary for linear invadosome formation and functionality (Figure 6).
In order to understand the invasive capacity of cells, we have also developed a 3D type I collagen plug assay (Figures 3 and 7). This assay allows us to determine the number of cells that have invaded into the collagen I plug, as well as the distance traveled, by using z-stack reconstructions.
<img alt="Figure 4" src="/files/ftp_upload/54911/54911fig4.jpg" /> 
Figure 4: Comparison of gelatin and gelatin-collagen I mixed matrices using confocal microscopy. Schematic representation showing the organization of fluorescent gelatin coverslips on the top panel. (A) Confocal z-stack reconstruction demonstrates that the fluorescent gelatin matrix forms a thin, uniform layer that covers the entire surface of the coverslip. (B) In the mixed matrix, after the deposition of fluorescent gelatin on the coverslips, type I collagen fibrils polymerize on top. The collagen I fibrils are stained in red. The collagen I matrix is thick and heterogeneous on the coverslip. The distribution of the collagen I fibers is dependent on the polymerization of collagen I &#945; chains. Scale bar = 10 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54911/54911fig5.jpg" /> 
Figure 5: Impact of gelatin and gelatin-collagen I matrices on invadosome formation and activity. The cells used for these assays are MDA-MB-231 breast cancer cells. (A) A schematic representation shows cells seeded on a fluorescent gelatin coverslip on the top panel. Degradation areas are visualized in black due to the reduction of fluorescence. Tks5 staining was used as an invadosome marker. On gelatin, the invadosomes that form are organized as dots. (B) A schematic representation shows cells seeded on a mixed matrix of gelatin and collagen I. Collagen I is labeled in red; the addition of type I collagen fibrils increases the ability of the cell to degrade gelatin. Interestingly, the invadosome marker Tks5 is reorganized, and the dots are replaced by linear structures representing linear invadosomes. Degradation areas are visualized in black due to the reduction of fluorescence. Scale bar = 5 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/54911/54911fig6.jpg" /> 
Figure 6: Confocal microscopy analysis of the molecular composition and organization of invadosomes in both gelatin and mixed-matrix conditions. (A) In the gelatin condition, invadosomes are organized in dots, and F-actin (red) co-localizes with Tks5 (green, bottom panel), but not with DDR1 (green, top panel). These are classical invadosomes. Scale bars = 5 &#181;m. (B) In the gelatin-collagen I mixed-matrix condition, DDR1 (green, top panel) colocalizes with the collagen I fibrils (red, top panel), Tks5 (green, bottom panel), and F-actin (red, bottom panel). The type I collagen fibrils induce the formation of linear invadosomes. Scale bars = 10 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/54911/54911fig7.jpg" /> 
Figure 7: Cell invasion assay of MDA-MB-231 cells in a collagen I plug. 1 h after seeding the cells, control inserts are fixed and stained. (A) z-stack acquisition of the collagen I plug is performed using confocal microscopy. The cells do not invade the collagen I plug at this time point; rather, they remain on top of the collagen I plug. Thus, this is used as the control time point when no invasion occurs. MMP activation is necessary for the cells to invade a collagen I plug at this density. (B) Three days after seeding, some cells invade the collagen I plug. This assay allows for the observation and quantification of cell invasion. In addition, it can be used to study the impact of various drugs or siRNAs, for example. Scale bar = 20 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54911/54911fig7large.jpg" target="_blank">Please click here to view a larger version of this figure. </a>

Watch this Scientific Journal Video about 2D and 3D Matrices to Study Linear Invadosome Formation and Activity at JoVE.com

2D and 3D Matrices to Study Linear Invadosome Formation and Activity

This protocol describes how to prepare a 2D mixed matrix, consisting of gelatin and collagen I, and a 3D collagen I plug to study linear invadosomes. These protocols allow for the study of linear invadosome formation, matrix degradation activity, and the invasion capabilities of primary cells and cancer cell lines.

Cell adhesion, migration, and invasion are involved in many physiological and pathological processes. For example, during metastasis formation, tumor ...

2d-and-3d-matrices-to-study-linear-invadosome-formation-and-activity

Nanyang Technological University

Research

JoVE Journal

Bioengineering

9.9K Views.  INSERM U1053. This protocol describes how to prepare a 2D mixed matrix, consisting of gelatin and collagen I, and a 3D collagen I plug to study linear invadosomes. These protocols allow for the study of linear invadosome formation, matrix degradation activity, and the invasion capabilities of primary cells and cancer cell lines. 

Video: 2D and 3D Matrices to Study Linear Invadosome Formation and Activity

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Electric Field-controlled Directed Migration of Neural Progenitor Cells in 2D and 3D Environments

Endogenous electric fields (EFs) occur naturally in vivo and play a critical role during tissue/organ development and regeneration, including that of the central nervous system1,2. These endogenous EFs are generated by cellular regulation of ionic transport combined with the electrical resistance of cells and tissues. It has been reported that applied EF treatment can promote functional repair of spinal cord injuries in animals and humans3,4. In particular, EF-directed cell migration has been demonstrated in a wide variety of cell types5,6, including neural progenitor cells (NPCs)7,8. Application of direct current (DC) EFs is not a commonly available technique in most laboratories. We have described detailed protocols for the application of DC EFs to cell and tissue cultures previously5,11. Here we present a video demonstration of standard methods based on a calculated field strength to set up 2D and 3D environments for NPCs, and to investigate cellular responses to EF stimulation in both single cell growth conditions in 2D, and the organotypic spinal cord slice in 3D. The spinal cordslice is an ideal recipient tissue for studying NPC ex vivo behaviours, post-transplantation, because the cytoarchitectonic tissue organization is well preserved within these cultures9,10. Additionally, this ex vivo model also allows procedures that are not technically feasible to track cells in vivo using time-lapse recording at the single cell level. It is critically essential to evaluate cell behaviours in not only a 2D environment, but also in a 3D organotypic condition which mimicks the in vivo environment. This system will allow high-resolution imaging using cover glass-based dishes in tissue or organ culture with 3D tracking of single cell migration in vitro and ex vivo and can be an intermediate step before moving onto in vivo paradigms.

This protocol demonstrates methods used to establish 2D and 3D environments in custom-designed electrotactic chambers, which can track cells in vivo/ex vivo using time-lapse recording at the single cell level, in order to investigate galvanotaxis/electrotaxis and other cellular responses to direct current (DC) electric fields (EFs).

Endogenous electric fields (EFs) occur naturally in vivo and play a critical role during tissue/organ development and regeneration, including that of the ...

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern tissue engineering. A technical bottleneck is the deposition of collagen in vitro, as it is notoriously slow, resulting in sub-optimal formation of connective tissue and subsequent tissue cohesion, particularly in skin models. Here, we describe a method which involves the addition of differentially-sized sucrose co-polymers to skin cultures to generate macromolecular crowding (MMC), which results in a dramatic enhancement of collagen deposition. Particularly, dermal fibroblasts deposited a significant amount of collagen I/IV/VII and fibronectin under MMC in comparison to controls.
The protocol also describes a method to decellularize crowded cell layers, exposing significant amounts of extracellular matrix (ECM) which were retained on the culture surface as evidenced by immunocytochemistry. Total matrix mass and distribution pattern was studied using interference reflection microscopy. Interestingly, fibroblasts, keratinocytes and co-cultures produced cell-derived matrices (CDM) of varying composition and morphology. CDM could be used as &#34;bio-scaffolds&#34; for secondary cell seeding, where the current use of coatings or scaffolds, typically from xenogenic animal sources, can be avoided, thus moving towards more clinically relevant applications.
In addition, this protocol describes the application of MMC during the submerged phase of a 3D-organotypic skin co-culture model which was sufficient to enhance ECM deposition in the dermo-epidermal junction (DEJ), in particular, collagen VII, the major component of anchoring fibrils. Electron microscopy confirmed the presence of anchoring fibrils in cultures developed with MMC, as compared to controls. This is significant as anchoring fibrils tether the dermis to the epidermis, hence, having a pre-formed mature DEJ may benefit skin graft recipients in terms of graft stability and overall wound healing. Furthermore, culture time was condensed from 5 weeks to 3 weeks to obtain a mature construct, when using MMC, reducing costs.

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

We present a protocol to obtain cell-derived matrices rich in extracellular matrix proteins, using macromolecular crowders (MMC). In addition, we present a protocol which incorporates MMC in 3D organotypic skin co-culture generation, which reduces culture time while maintaining maturity of construct.

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic significance. Possible reasons for the lack of exploration are the experimental limitations and technical challenges that render the study of cell division in 3D culture inefficient. Here, we describe an imaging-based method to efficiently study mammalian cell division and cell-matrix interactions in 3D collagen matrices. Cells labeled with fluorescent H2B are synchronized using the combination of thymidine blocking and nocodazole treatment, followed by a mechanical shake-off technique. Synchronized cells are then embedded into a 3D collagen matrix. Cell division is monitored using live-cell microscopy. The deformation of collagen fibers during and after cell division, which is an indicator of cell-matrix interaction, can be monitored and quantified using quantitative confocal reflection microscopy. The method provides an efficient and general approach to study mammalian cell division and cell-matrix interactions in a physiologically relevant 3D environment. This approach not only provides novel insights into the molecular basis of the development of normal tissue and diseases, but also allows for the design of novel diagnostic and therapeutic approaches.

This protocol efficiently studies mammalian cell division in 3D collagen matrices by integrating synchronization of cell division, monitoring of division events in 3D matrices using live-cell imaging technique, time-resolved confocal reflection microscopy and quantitative imaging analysis.

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic ...

Bioprinting of Cartilage and Skin Tissue Analogs Utilizing a Novel Passive Mixing Unit Technique for Bioink Precellularization

Bioprinting is a powerful technique for the rapid and reproducible fabrication of constructs for tissue engineering applications. In this study, both cartilage and skin analogs were fabricated after bioink pre-cellularization utilizing a novel passive mixing unit technique. This technique was developed with the aim to simplify the steps involved in the mixing of a cell suspension into a highly viscous bioink. The resolution of filaments deposited through bioprinting necessitates the assurance of uniformity in cell distribution prior to printing to avoid the deposition of regions without cells or retention of large cell clumps that can clog the needle. We demonstrate the ability to rapidly blend a cell suspension with a bioink prior to bioprinting of both cartilage and skin analogs. Both tissue analogs could be cultured for up to 4 weeks. Histological analysis demonstrated both cell viability and deposition of tissue specific extracellular matrix (ECM) markers such as glycosaminoglycans (GAGs) and collagen I respectively.

Cartilage and skin analogs were bioprinted using a nanocellulose-alginate based bioink. The bioinks were cellularized prior to printing via a single step passive mixing unit. The constructs were demonstrated to be uniformly cellularized, have high viability, and exhibit favorable markers of differentiation.

Bioprinting is a powerful technique for the rapid and reproducible fabrication of constructs for tissue engineering applications. In this study, both ...

Bioprintable Alginate/Gelatin Hydrogel 3D In Vitro Model Systems Induce Cell Spheroid Formation

The cellular, biochemical, and biophysical heterogeneity of the native tumor microenvironment is not recapitulated by growing immortalized cancer cell lines using conventional two-dimensional (2D) cell culture. These challenges can be overcome by using bioprinting techniques to build heterogeneous three-dimensional (3D) tumor models whereby different types of cells are embedded. Alginate and gelatin are two of the most common biomaterials employed in bioprinting due to their biocompatibility, biomimicry, and mechanical properties. By combining the two polymers, we achieved a bioprintable composite hydrogel with similarities to the microscopic architecture of a native tumor stroma. We studied the printability of the composite hydrogel via rheology and obtained the optimal printing window. Breast cancer cells and fibroblasts were embedded in the hydrogels and printed to form a 3D model mimicking the in vivo microenvironment. The bioprinted heterogeneous model achieves a high viability for long-term cell culture (&#62; 30 days) and promotes the self-assembly of breast cancer cells into multicellular tumor spheroids (MCTS). We observed the migration and interaction of the cancer-associated fibroblast cells (CAFs) with the MCTS in this model. By using bioprinted cell culture platforms as co-culture systems, it offers a unique tool to study the dependence of tumorigenesis on the stroma composition. This technique features a high-throughput, low cost, and high reproducibility, and it can also provide an alternative model to conventional cell monolayer cultures and animal tumor models to study cancer biology.

Bioprintable Alginate/Gelatin Hydrogel 3D In Vitro Model Systems Induce Cell Spheroid Formation

We developed a heterogeneous breast cancer model consisting of immortalized tumor and fibroblast cells embedded in a bioprintable alginate/gelatin bioink. The model recapitulates the in vivo tumor microenvironment and facilitates the formation of multicellular tumor spheroids, yielding insight into the mechanisms driving tumorigenesis.

The cellular, biochemical, and biophysical heterogeneity of the native tumor microenvironment is not recapitulated by growing immortalized cancer cell ...

Novel Process for 3D Printing Decellularized Matrices

3D bioprinting aims to create custom scaffolds that are biologically active and accommodate the desired size and geometry. A thermoplastic backbone can provide mechanical stability similar to native tissue while biologic agents offer compositional cues to progenitor cells, leading to their migration, proliferation, and differentiation to reconstitute the original tissues/organs1,2. Unfortunately, many 3D printing compatible, bioresorbable polymers (such as polylactic acid, PLA) are printed at temperatures of 210 &#176;C or higher - temperatures that are detrimental to biologics. On the other hand, polycaprolactone (PCL), a different type of polyester, is a bioresorbable, 3D printable material that has a gentler printing temperature of 65 &#176;C. Therefore, it was hypothesized that decellularized extracellular matrix (DM) contained within a thermally protective PLA barrier could be printed within PCL filament and remain in its functional conformation. In this work, osteochondral repair was the application for which the hypothesis was tested. As such, porcine cartilage was decellularized and encapsulated in polylactic acid (PLA) microspheres which were then extruded with polycaprolactone (PCL) into filament to produce 3D constructs via fused deposition modeling. The constructs with or without the microspheres (PLA-DM/PCL and PCL(-), respectively) were evaluated for differences in surface features.

This protocol describes the production of polycaprolactone (PCL) filament with embedded polylactic acid (PLA) microspheres which contain decellularized matrices (DM) for 3D printing of structural tissue engineering constructs.

3D bioprinting aims to create custom scaffolds that are biologically active and accommodate the desired size and geometry. A thermoplastic backbone can ...

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

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

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

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

Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current in vitro screening models that have sufficient throughput only have limited physiological relevance, which hinders the translation of findings from in vitro to in vivo. On the other hand, microfluidic cell culture platforms have shown unparalleled physiological relevancy in vitro, but often lack the required throughput, scalability and standardization. We demonstrate a robust platform to study angiogenesis of endothelial cells derived from human induced pluripotent stem cells (iPSC-ECs) in a physiological relevant cellular microenvironment, including perfusion and gradients. The iPSC-ECs are cultured as 40 perfused 3D microvessels against a patterned collagen-1 scaffold. Upon the application of a gradient of angiogenic factors, important hallmarks of angiogenesis can be studied, including the differentiation into tip- and stalk cell and the formation of perfusable lumen. Perfusion with fluorescent tracer dyes enables the study of permeability during and after anastomosis of the angiogenic sprouts. In conclusion, this method shows the feasibility of iPSC-derived ECs in a standardized and scalable 3D angiogenic assay that combines physiological relevant culture conditions in a platform that has the required robustness and scalability to be integrated within the drug screening infrastructure.

This method describes the culture of iPSC-derived endothelial cells as 40 perfused 3D microvessels in a standardized microfluidic platform. This platform enables the study of gradient-driven angiogenic sprouting in 3D, including anastomosis and stabilization of the angiogenic sprouts in a scalable and high-throughput manner.

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current ...