The authors thank W. Sun and K. Jansen for the critical discussions, and acknowledge support by the Nano Biomechanics Lab at the National University of Singapore. N.A.K. acknowledges support by a Marie Curie IIF Fellowship.

1. Cell Harvesting
<ol>
	<li>Obtain MD-MBA-231 cells from the 37 &#176;C, 5% CO2 incubator. Detach cells from tissue culture plate using 0.5% Trypsin-EDTA solution. Use 1 ml of Trypsin-EDTA solution for cell cultured in a T25 flask.</li>
	<li>Pellet cells in a 15 ml conical tube by centrifugation at 200 &#215; g for 4 min, aspirate the supernatant, and resuspend cells in 5 ml of culture media.</li>
	<li>Count cell density, &#961;, using a hemocytometer. 
	Note: To prepare the cell-seeded inner gel, the cell suspension will later on be diluted 10&#215; to reach the final cell seeding density. Therefore, 10&#215; concentrated cell suspension is required.</li>
	<li>Calculate the volume of medium required to achieve a 10&#215; cell concentration: 
	<img alt="Equation 1" fo:content-width="2in" src="/files/ftp_upload/52735/52735eq1.jpg" width="175" /> 
	Note: A final cell seeding density, cfinal, of around 2 &#215; 106 cells/ml is recommended for MD-MBA-231 cells and is used in this protocol. Other seeding densities can also be explored for other cell types.</li>
	<li>Pellet cells one more time in a 15 ml conical tube by centrifugation at 200 &#215; g for 4 min, and aspirate the supernatant.</li>
	<li>Resuspend the cells in the required amount (Vmedium calculated in step 1.3) of serum-free cell culture medium thoroughly to minimize cell clumping. 
	Note: Phenol red is auto-fluorescent, and can interfere with fluorescence/reflectance imaging. Use of phenol red-free medium may be considered to achieve best image quality.</li>
</ol>
2. Preparation of Collagen Solutions
<ol>
	<li>Obtain the stock collagen solution, 10&#215; PBS buffer, Milli-Q H2O, 0.1 M NaOH, and several microcentrifuge tubes. Keep all on ice to prevent premature collagen polymerization, and maintain sterile condition.</li>
	<li>Equilibrate a sterile glass-bottom dish by pre-warming it in the 37 &#176;C incubator. 
	Note: All volumes in this protocol have been optimized for a glass-bottom dish with 12 mm well. If other dish type is used, adjust the volumes correspondingly.</li>
	<li>Calculate the required volume needed to prepare 50 &#181;l of 2.4 mg/ml collagen solution for the inner gel (solution I) based on the collagen stock concentration. 
	Note: Other collagen concentrations for the inner gel can also be used.</li>
	<li>In a sterile environment (typically a biosafety hood), slowly add 5 &#181;l of 10&#215; PBS buffer to the required amount of collagen stock solution (calculated in step 2.3) with gentle swirling. Take care to avoid air bubble formation.</li>
	<li>Adjust the pH of the mixture to 7.4 using 0.1 M NaOH using calibrated pH-meter. As a rough guide, use about 5 &#181;l to bring the pH close to 7.4 (the amount varies depending on the stock concentration and pH). 
	Note: Note that the volumes involved in this step is too small for the use of standard pH meter. Use one of the following tricks:
	<ol>
		<li>Prepare collagen solutions for multiple samples. Adjust the pH in bulk using standard pH meter and distribute the collagen solutions across the samples.</li>
		<li>Alternatively, adjust the pH of a collagen solution in larger volume (i.e., volume that allow the use of standard pH meter). Note the amount of NaOH needed to bring the pH to the final pH. Scale down the volumes and use the appropriate amount of NaOH for the experiment. Confirm the pH value using Litmus paper.</li>
		<li>Otherwise, use a micro pH electrode to more accurately adjust the pH of small quantities.</li>
	</ol>
	</li>
	<li>Bring the solution to a volume of 45 &#181;l using H2O. Perform all steps on ice to prevent premature collagen polymerization.</li>
</ol>
3. Formation of Concentric Gel Culture
<ol>
	<li>Take the pre-warmed glass-bottom dish (see step 2.2) from the incubator.</li>
	<li>Add 5 &#181;l of the 10&#215; concentrated cell suspension (prepared in step 1.5) to solution I. Resuspend thoroughly. Take care to avoid air bubble formation. The mixture now has a volume of 50 &#181;l and contains the final collagen concentration (2.4 mg/ml) and cell density (cfinal = 2 &#215; 106 cells/ml).</li>
	<li>Add 20 &#181;l of the cell-containing solution I slowly to the center of the well, so that it forms a dome-shaped droplet (Figure 1A). Take care to avoid bubble formation at this step. If a bubble forms, carefully but quickly try to either rupture it or suck it out using a pipette tip. Gently place the dish back in the incubator to let the inner gel polymerize for 45 min.</li>
	<li>Prepare solution O (for the outer, acellular collagen gel) about 15 min before the end of this incubation step. 
	Note: The outer gel condition can be varied in terms of collagen concentration and polymerization pH to obtain networks with different microstructures10. In this protocol, focus on a 1.5 &#8211; 4.0 mg/ml collagen gel polymerized at a pH of 7.4.
	<ol>
		<li>Based on the collagen stock concentration, calculate the required volume needed to prepare 200 &#181;l of collagen solution O at the final concentration.</li>
	</ol>
	</li>
	<li>Add 20 &#181;l of 10&#215; PBS buffer slowly to the required amount of collagen stock solution (calculated in step 3.4) with gentle swirling. Adjust the pH of the mixture to the final pH using 0.1 M NaOH with the use of calibrated pH-meter. See note to step 2.5 regarding pH adjustment.</li>
	<li>Bring the solution to a final volume of 200 &#181;l using H2O. Perform all steps on ice to prevent premature collagen polymerization.</li>
	<li>Take the dish from the incubator after 45 min polymerization of the inner gel (see step 3.3). Gently add 180 &#181;l of solution O on top of the inner gel, so that the solution completely covers the inner gel and fills up the well (Figure 1B).
	<ol>
		<li>Perform this step carefully without stirring the solution, which can lead to non-uniform fiber orientations in the outer gel. Take care to avoid touching the inner gel with a pipette tip, and to avoid the formation of bubbles or air pockets. If a bubble forms, carefully but quickly try to either rupture it or suck it out using a pipette tip. Gently place the dish back in the incubator to let the outer gel polymerize.</li>
	</ol>
	</li>
	<li>Take the dish from the incubator after 45 min of polymerization. The gel should already be fairly solidified at this point, although it can still detach from the bottom surface if handled roughly.
	<ol>
		<li>Gently pour 2 ml of warmed cell culture medium to the dish (Figure 1C). Make sure that the gel is completely submerged in the medium. Refresh the medium every 2 &#8211; 3 days throughout the duration of culture.</li>
	</ol>
	</li>
</ol>
4. Live-cell Imaging
<ol>
	<li>Perform imaging using an inverted confocal microscope equipped with long-term live-cell imaging capability. Include a built-in incubation chamber with a temperature (37 &#176;C) and CO2 (5%) control. Switch on the microscope and warm up the stage at least 1 hr before starting the experiment. 
	Note: Use objective lens with long working distance to optimize the observation and localization of cells in the 3D gels.</li>
	<li>Incubate the gel in medium containing 5 &#181;l of fluorescent cell tracker dye for 30 min to allow accurate localization of the cells in the 3D system. Subsequently, remove unbound dye by washing three times with 1&#215; PBS. Afterwards, add cell culture medium to the dish.</li>
	<li>Take the dish from the incubator and place it on the microscope stage (Figure 1D). 
	Note: Live cell imaging can start in principle right after the polymerization of the outer gel. However at this point, the cells in the inner gel have not spread yet. To examine the migration of the first cells that have invaded the outer gel, live-cell imaging can start 24 hr (depending on the cell type) after the initiation of culture. As a rough guide, around 12 - 14 days of culture are needed for most of the cells in the inner gel to enter the outer gel.</li>
	<li>Select Volumes of View (VoV&#8217;s) in regions in the outer gel surrounding the inner gel. After 24 hr of incubation, the cell population will have spread, crossed the interface between the inner and outer gels, and started to invade the outer gel.
	<ol>
		<li>For the VoV&#8217;s, include the gel regions immediately next to the gel interface, intermediate regions, and regions close to the outskirt of the outer gel7. Exclude regions closer than 50 &#181;m from the bottom and side surfaces, as well as from the top of the gel, to avoid possible edge effects. Each VoV typically measures 647 &#215; 647 &#215; 100 &#181;m3 (in x, y, and z directions, respectively), with 5 &#181;m interval in the z-stack.</li>
	</ol>
	</li>
	<li>Ensure imaging modes, channels/filters, exposure times, and image resolutions are correctly selected. For label-free imaging of the collagen network, use confocal reflectance microscopy simultaneously during the time-lapse live-cell imaging.</li>
	<li>Take sample images, plot the intensity histograms, and adjust the gains and offsets to observe sufficient signal and avoid saturation by ensuring that the histogram lies between zero and the maximum intensity. Do not change these settings any more throughout the duration of the experiment.</li>
	<li>Take time-lapse images of the selected VoV&#8217;s, with a time interval, &#916;t, of 10 min for 8 hrs (or longer if necessary).</li>
</ol>
5. Cell Tracking and Data Analysis
<ol>
	<li>Carry out quantitative image post-processing from the z-stack images using appropriate image processing software.
	<ol>
		<li>Segment the time-lapse images to automatically select the 3D cell positions in (x,y,z,t).</li>
		<li>For every frame, manually check the localization accuracy and remove false positives due to cell debris and cellular protrusions that may have been mistaken for cells. Remove proliferative cells from the analysis and split overlapping or connected cells into distinct objects.</li>
	</ol>
	</li>
	<li>Generate 3D time-lapse cell tracks from the cell coordinates (x,y,z,t) obtained in the previous step by linking the location of each cell in the time sequence.</li>
	<li>Eliminate random and system noise by removing tracks shorter than a threshold track length (typically 20 min).</li>
	<li>Correct for sample drift by subtracting the overall net displacements from the tracks if necessary.</li>
	<li>Calculate cell displacement, <img alt="Equation 2" fo:content-width="0.8in" src="/files/ftp_upload/52735/52735eq2.jpg" width="80" />, and cell migration distance, <img alt="Equation 3" fo:content-width="1in" src="/files/ftp_upload/52735/52735eq3.jpg" width="100" />, from the observed cell trajectories, where is a vector representing the 3D location of a cell at time and is the total number of time points.
	<ol>
		<li>Calculate cell speed as S = d/(n &#8226; &#916;t), where &#916;t is the time interval between frames. Calculate cell migration directionality (or persistence) using P = &#916;d/d. This simple measure of persistence implies that for P = 0 the net displacement is zero and for P = 1 the trajectory is a straight directional line.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

In this protocol we describe an in vitro assay to study the 3D migrational behavior of cells in matrix environments that topologically resemble ECMs encountered in vivo. There are three main strengths of this assay as compared to other currently available methods. First, this assay allows one to simultaneously examine the cell migration mechanisms at both population level and individual cell level. This opens up possibilities of studying collective cell migration13, which has to date been lar...

Migration of cells plays a key role in various physiological responses such as embryonic development, haemostasis, and immune response as well as in pathological processes such as vascular diseases, inflammation, and cancer1. Dissecting the biochemical and biophysical factors underlying cell migration is therefore fundamentally important not only to understand the basic principles of cellular functions, but also to advance various biomedical applications, such as in tissue engineering, anti-metastasis and anti-inflammatory drug development. Since in vivo observation is technically challenging, a lot of efforts has been focused on in vitro recapitulation of cell migration.
In vitro methods to study cell migration have largely been designed for assays on two dimensional (2D) surfaces, most notably the scratch or wound healing assay2. Such assays offer simple experimental setup, easy live-cell imaging, and provide useful insights into various biochemical mechanisms underlying cell migration. However, these assays do not account for extracellular matrix (ECM) architecture and remodeling, which are critical aspects in understanding in vivo migration. Recently, it has been increasingly appreciated that a 3D culture model, often in collagen-based matrices3, provides a platform that better resembles the in vivo situation. Indeed, cells exhibit migrational dynamics that are distinct from those on 2D surfaces, especially due to the different dimensionality of the environment4. Moreover, the biophysical and mechanical properties of the matrix sensitively affect cell migration5, including in the context of tumor cell invasion6.
Here, we present a method to study 3D cell migration behavior in ECM with biophysical properties that can be easily varied with preparation conditions. The cells are seeded in an &#8220;inner gel&#8221; and are allowed to escape into and invade the initially acellular &#8220;outer gel&#8221;. The method relies on the cell&#8217;s ability to recognize the presence of, and propensity to migrate into, cell-free regions in the outer gel, which is closely linked to cell mechanosensing7. In this study, we employ collagen networks as the ECMs invaded by highly invasive breast cancer cells, MD-MBA-231. The mechanical properties and microstructure of both the inner and outer gels can be tuned8 and characterized9 to achieve physiologically relevant conditions. Reconstruction and analysis of the cell tracks allow detailed quantitative examination of the spatiotemporal migration behavior at both population level and individual cell level. Importantly, the setup of the concentric gel system mimics the in vivo tissue topology faced by migrating cells, especially invading cancer cells, thus offering important insights into the physical mechanisms of cell migration and metastasis.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Cell culture incubator</td>
			<td>Fisher Scientific Pte Ltd</td>
			<td>Model: 371, S/No 318854-6055</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Nikon A1R</td>
			<td></td>
			<td>Inverted confocal laser scanning microscope equipped with incubator chamber</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM)</td>
			<td>Life Technologies</td>
			<td>11965-092</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Life Technologies</td>
			<td>10082-147</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent CellTracker dye CMTMR</td>
			<td>Life Technologies</td>
			<td>C2927</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass-bottom dish</td>
			<td>IWAKI Cell Biology</td>
			<td>3931-035</td>
			<td>35 mm diameter dish with 12 mm diameter glass-bottom well</td>
		</tr>
		<tr>
			<td>Hemocytometer</td>
			<td>iN CYTO</td>
			<td>DHC-N01 (Neubauer Improved)</td>
			<td></td>
		</tr>
		<tr>
			<td>Microprocessor pH meter</td>
			<td>Hanna Instruments</td>
			<td>pH 211</td>
			<td></td>
		</tr>
		<tr>
			<td>Nutragen Collagen</td>
			<td>Advanced BioMatrix</td>
			<td>#5010-D</td>
			<td>Acid-solubilized bovine collagen type I (stock pH ~ 2)</td>
		</tr>
		<tr>
			<td>Objective lens</td>
			<td>Nikon</td>
			<td></td>
			<td>CFI Super Plan Fluor ELWD ADM 20XC, W.D. 8.2-6.9mm, NA 0.45.</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Life Technologies</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Sartorius</td>
			<td>S/No 29153352</td>
			<td>Basic pH Meter PB-11</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA</td>
			<td>Life Technologies</td>
			<td>15400-054</td>
			<td></td>
		</tr>
	</tbody>
</table>

concentric gel system to study the biophysical role of matrix microenvironment on 3d cell migration

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to tumor and cancer metastasis. Despite intense research efforts, the basic biochemical and biophysical principles of cell migration are still not fully understood, especially in the physiologically relevant three-dimensional (3D) microenvironments. Here, we describe an in vitro assay designed to allow quantitative examination of 3D cell migration behaviors. The method exploits the cell&#8217;s mechanosensing ability and propensity to migrate into previously unoccupied extracellular matrix (ECM). We use the invasion of highly invasive breast cancer cells, MDA-MB-231, in collagen gels as a model system. The spread of cell population and the migration dynamics of individual cells over weeks of culture can be monitored using live-cell imaging and analyzed to extract spatiotemporally-resolved data. Furthermore, the method is easily adaptable for diverse extracellular matrices, thus offering a simple yet powerful way to investigate the role of biophysical factors in the microenvironment on cell migration.

The concentric gel assay presented here was performed using highly invasive breast cancer cells, MDA-MB-231, with 2.4 mg/ml inner collagen gel and a cell seeding density of = 2 &#215; 106 cells/ml, as an example. As shown in Figure 2, typically after a few days of culture, the cells breached the inner&#8211;outer gel interface and started to invade the outer gel. The cell population spread predominantly radially outwards.
The polymerization conditions of the outer g...

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Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

The mechanical properties and microstructure of the extracellular matrix strongly affect 3D migration of cells. An in vitro method to study the spatiotemporal cell migration behavior in biophysically variable environments, at both population and individual cell levels, is described.

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to ...

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Research

JoVE Journal

Bioengineering

8.5K 조회수.  FOM Institute AMOLF.  기계적 특성 및 세포 외 매트릭스의 미세 강하게 세포의 3 차원 이동에 영향을 미칩니다. 시험 관내 방법은 개별 세포 집단 수준에서 모두 biophysically 변수 환경에서 시공간 세포 이동 동작을 연구하기 위해, 설명한다. 

비디오: Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

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 ...

Study of Cell Migration in Microfabricated Channels

The method described here allows the study of cell migration under confinement in one dimension. It is based on the use of microfabricated channels, which impose a polarized phenotype to cells by physical constraints. Once inside channels, cells have only two possibilities: move forward or backward. This simplified migration in which directionality is restricted facilitates the automatic tracking of cells and the extraction of quantitative parameters to describe cell movement. These parameters include cell velocity, changes in direction, and pauses during motion. Microchannels are also compatible with the use of fluorescent markers and are therefore suitable to study localization of intracellular organelles and structures during cell migration at high resolution. Finally, the surface of the channels can be functionalized with different substrates, allowing the control of the adhesive properties of the channels or the study of haptotaxis. In summary, the system here described is intended to analyze the migration of large cell numbers in conditions in which both the geometry and the biochemical nature of the environment are controlled, facilitating the normalization and reproducibility of independent experiments.

A quantitative method to study spontaneous migration of cells in a one-dimensional confined microenvironment is described. This method takes advantage of microfabricated channels and can be used to study migration of large number of cells under different conditions in single experiments.

The method described here allows the study of cell migration under confinement in one dimension. It is based on the use of microfabricated channels, which ...

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic development, wound healing, and immune responses. However, cell migration is also a key mechanism in cancer enabling these cancer cells to detach from the primary tumor to start metastatic spreading. Within the past years various cell migration assays have been developed to analyze the migratory behavior of different cell types. Because the locomotory behavior of cells markedly differs between a two-dimensional (2D) and three-dimensional (3D) environment it can be assumed that the analysis of the migration of cells that are embedded within a 3D environment would yield in more significant cell migration data. The advantage of the described 3D collagen matrix migration assay is that cells are embedded within a physiological 3D network of collagen fibers representing the major component of the extracellular matrix. Due to time-lapse video microscopy real cell migration is measured allowing the determination of several migration parameters as well as their alterations in response to pro-migratory factors or inhibitors. Various cell types could be analyzed using this technique, including lymphocytes/leukocytes, stem cells, and tumor cells. Likewise, also cell clusters or spheroids could be embedded within the collagen matrix concomitant with analysis of the emigration of single cells from the cell cluster/ spheroid into the collagen lattice. We conclude that the 3D collagen matrix migration assay is a versatile method to analyze the migration of cells within a physiological-like 3D environment.

Cell migration is a biological phenomenon that is involved in a plethora of physiological, such as wound healing and immune responses, and pathophysiological processes, like cancer. The 3D-collagen matrix migration assay is a versatile tool to analyze the migratory properties of different cell types within in a 3D physiological-like environment.

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic ...

Planar Gradient Diffusion System to Investigate Chemotaxis in a 3D Collagen Matrix

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to their surrounding area, leading to cell movement and migration. In order to understand the mechanisms that can alter and/or affect cell migration, one can study these forces. In theory, understanding the fundamental mechanisms and forces underlying cell migration holds the promise of effective approaches for treating diseases and promoting cellular transplantation. Unfortunately, modern chemotaxis chambers that have been developed are usually restricted to two dimensions (2D) and have complex diffusion gradients that make the experiment difficult to interpret. To this end, we have developed, and describe in this paper, a direct-viewing chamber for chemotaxis studies, which allows one to overcome modern chemotaxis chamber obstacles able to measure cell forces and specific concentration within the chamber in a 3D environment to study cell 3D migration. More compelling, this approach allows one to successfully model diffusion through 3D collagen matrices and calculate the coefficient of diffusion of a chemoattractant through multiple different concentrations of collagen, while keeping the system simple and user friendly for traction force microscopy (TFM) and digital volume correlation (DVC) analysis.

Cell migration is an important part of human development and life. In order to understand the mechanisms that can alter cell migration, we present a planar gradient diffusion system to investigate chemotaxis in a 3D collagen matrix, which allows one to overcome modern diffusion chamber limitations of existing assays.

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to ...

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 ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

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

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

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

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

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

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

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

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

Labeling of Extracellular Vesicles for Monitoring Migration and Uptake in Cartilage Explants

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their cellular source, such as platelet lysate (PL). When used as treatment, EVs are expected to enter the target cells and effect a response from these. In this research, PL-derived EVs have been studied as a cell-free treatment for osteoarthritis (OA). Thus, a method was set up to label EVs and test their uptake on cartilage explants. PL-derived EVs are labeled with the lipophilic dye PKH26, washed twice through a column, and then tested in an in vitro inflammation-driven&#160;OA model for 5 h after particle quantification by nanoparticle tracking analysis (NTA). Hourly, cartilage explants are fixed, paraffined, cut into 6 &#181;m sections to mount on slides, and observed under a confocal microscope. This allows verification of whether EVs enter the target cells (chondrocytes) during this period and analyze their direct effect.

Here, we present a protocol to label platelet lysate-derived extracellular vesicles to monitor their migration and uptake in cartilage explants used as a model for osteoarthritis.

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their ...

Microgel-Extracellular Matrix Composite Support for the Embedded 3D Printing of Human Neural Constructs

The embedded 3D printing of cells inside a granular support medium has emerged in the past decade as a powerful approach for the freeform biofabrication of soft tissue constructs. However, granular gel formulations have been restricted to a limited number of biomaterials that allow for the cost-effective generation of large amounts of hydrogel microparticles. Therefore, granular gel support media have generally lacked the cell-adhesive and cell-instructive functions found in the native extracellular matrix (ECM).
To address this, a methodology has been developed for the generation of self-healing annealable particle-extracellular matrix (SHAPE) composites. SHAPE composites consist of a granular phase (microgels) and a continuous phase (viscous ECM solution) that, together, allow for both programmable high-fidelity printing and an adjustable biofunctional extracellular environment. This work describes how the developed methodology can be utilized for the precise biofabrication of human neural constructs.
First, alginate microparticles, which serve as the granular component in the SHAPE composites, are fabricated and combined with a collagen-based continuous component. Then, human neural stem cells are printed inside the support material, followed by the annealing of the support. The printed constructs can be maintained for weeks to allow the differentiation of the printed cells into neurons. Simultaneously, the collagen continuous phase allows for axonal outgrowth and the interconnection of regions. Finally, this works provides information on how to perform live-cell fluorescence imaging and immunocytochemistry to characterize the 3D-printed human neural constructs.

This work describes a protocol for the freeform embedded 3D printing of neural stem cells inside self-healing annealable particle-extracellular matrix composites. The protocol enables the programmable patterning of interconnected human neural tissue constructs with high fidelity.

The embedded 3D printing of cells inside a granular support medium has emerged in the past decade as a powerful approach for the freeform biofabrication ...