Funding for this work was partially supported through the US National Institutes of Health High Priority, Short-Term Project Award (1R56DK126020-01) and a philanthropic gift from the Kurtin Trust. J.T. was funded by the National Science Foundation Graduate Research Fellowship.&#160;Figure schematics created with BioRender.com.

The details of the reagents and the equipment used in the study are listed in the Table of Materials.
1. Granular hydrogel preparation
NOTE: MAP particles used throughout this protocol are 3.2 wt% w/v gel with 45.88 mg/mL PEG-MAL (10 kDa), 0.82 mg/mL RGD, 8.06 mg/mL MethMal19, and 5.62 mg/mL MMP-2 degradable crosslinker. The mechanical stiffness of the gel is 15-20 kPa to match dermal stiffness17.
<ol>
	<li>Generate granular hydrogel particles and prepare for cell culture as normal. 
	NOTE: This protocol describes the sterile preparation of Microporous Annealed Particle gel, the production of which is detailed by Roosa et al.18.</li>
	<li>Prepare the granular hydrogel particles for in vitro usage by sterilizing with three washes of 70% isopropyl alcohol followed by three washes of sterile 1x PBS.</li>
	<li>Prepare a sterile 0.2 mM lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) in media solution by dissolving LAP powder in ultrapure water and passing the solution through a 0.22 &#181;m sterile filter. Add this solution 1:1 v/v with the amount of gel necessary to conduct the experiment.</li>
	<li>Incubate the 0.2 mM LAP, gel, and media solution at 37 &#176;C on a tube rotator at 20 rpm for at least 30 min prior to proceeding to allow diffusion of LAP throughout the microparticles.</li>
	<li>Once 30 min have elapsed, centrifuge the particle suspension at 18,000 x g for 5 min at 25 &#176;C. Aspirate the supernatant. 
	NOTE: Overall dryness of MAP particles can vary due to differences in chemistry, hydrophilicity, and particle size. For consistency, it is best practice to centrifuge the particle suspension as described, mix the particles in the tube using a positive displacement pipette, and then repeat the centrifugation step.</li>
</ol>
2. Monolayer-based Ascending Migration Assay (MAMA) method: Cell culture and imaging
NOTE: Cells were imaged using the FITC channel (488 nm). The dye used for the cells had an excitation at 492 nm and an emission at 517 nm. 10x magnification provides increased detail over 4x magnification, but either can be used.
<ol>
	<li>Thaw human dermal fibroblasts (HDFs) according to the manufacturer&#39;s protocol. Passage as needed until the desired passage is achieved; generally, primary cells maintain genetic and phenotypic makeup through P5.</li>
	<li>Plate 120,000 cells/cm2 for at least n = 6 in a 96-well plate or desired plate size using the recommended media volume per well for suspension, gently aspirating before each addition. Let cells attach overnight, as shown in Figure 1A, which will result in approximately 80% confluency the next day. 
	NOTE: Generally, center column and row wells are best for optimal gel spreading. The authors use up to 24 wells for a 96-well plate (rows B-G and columns 5-8).</li>
	<li>Prepare gel conditions as noted above, and while incubating in LAP, remove the media with an aspirator or a pipette from the well plate of cells. Be careful not to disturb the bottom of the plate.</li>
	<li>Add cell tracking dye to wells according to manufacturer instructions, represented in Figure 1B. Ensure the gel is completely prepared as described in step 1 before aspirating cell tracking dye from wells.</li>
	<li>Add 20 &#181;L of each gel condition to the wells with a positive displacement pipette without touching the bottom of the plate. 
	NOTE: Best assay conditions occur when the gel is pipetted directly to the center of the well.</li>
	<li>Using a plate-spinning rotor centrifuge attachment at 25 &#176;C, spin at 100 x g for 15 s with acceleration and deceleration of 8 to flatten the gel. Flip the plate 180&#176; and spin again at 100 x g for 15 s to ensure even gel distribution across the well bottom, as seen in Figure 1C.</li>
	<li>Aseptically photo-crosslink the gel from the top (Figure 1D) by applying focused light (365 nm, 34.4 mW/cm2) to the sample for 30 s to anneal the scaffold, adding 200 &#181;L of media to each well of cells after all scaffolds are formed. Let the cells incubate at 37 &#176;C for 30 min to allow them to attach to the granular scaffold before imaging.</li>
	<li>To capture migration behavior summarized in Figure 1E, image cells using a confocal microscope. Find the lowest point of focus for an area of the plate where cells are at least 80% confluent and set it as the lower edge of the z-stack.
	<ol>
		<li>Find the highest point of the cell fluorescent signal and set it as the upper edge of the z-stack. Use a 5 &#181;m or smaller step size for best resolution.</li>
	</ol>
	</li>
	<li>Image at least three wells to represent the cell monolayer and non-migrating cell heights. The t = 0 timepoint is shown from both the top view and side view in Figure 2A. Incubate overnight after imaging is complete. 
	NOTE: These cells were imaged with a 4x objective, and exposure was kept consistent for all wells.</li>
	<li>At t = 24 h, cells will begin to ascend through the granular scaffold, as seen in Figure 2B from top and side views . Repeat imaging steps as performed for t = 0 h using the same parameters. Use the previous height of the stage as a reference or find any cells that have not yet migrated and set that as the lower edge of the z-stack.</li>
	<li>Repeat steps for all wells, ensuring each well is saved as a separate image for ease of analysis. 
	NOTE: Timepoints may be imaged for longer timepoints depending on experimental constraints and method of cell tracking fluorescence.</li>
	<li>Scaffolds may be fixed and stained for additional metrics. At the desired end timepoint, aspirate the media from wells with a pipette and discard. Gently wash each well with 200 &#181;L of sterile PBS twice for 5 min each. Add 200 &#181;L of 4% Paraformaldehyde (PFA) for 20 min, then aspirate and discard. Wells can be stained immediately or stored at 4 &#176;C in 1x PBS for up to a week.</li>
</ol>
3. Monolayer-based Ascending Migration Assay (MAMA) method: 3D image analysis
<ol>
	<li>For batch conversions, open the IMARIS Image Conversion software. Drag and drop the microscopy images into the conversion software and choose a folder inside the software Arena to import. Press Start All. The converted images will appear in the Arena as .ims files. 
	NOTE: If voxel size is not included in the image metadata, refer to individual confocal imaging specifications to find the value. Alternatively, voxel size can be approximated as the step size used during imaging.</li>
	<li>Open the software by double-clicking the desktop IMARIS Arena icon and selecting an image from the Arena.</li>
	<li>The image is automatically loaded into the &#39;3D View&#39; analysis tab seen in the toolbar icon panel at the top. Click on the Image Proc tab in the main toolbar.</li>
	<li>In the top left of the side panel, click on the dropdown menu for Channel 1 and select Background Subtraction. Press Ok at the bottom of the panel to return to &#39;3D View&#39;. Representative images for t = 0 h and t = 24 h are in Figure 2A,B, respectively.</li>
	<li>In the small toolbar just above the side panel menu, click on the icon with rounded blue shapes, Add new Surfaces, to create an editable objects tab named &#39;Surfaces 1&#39;.</li>
	<li>An interface for creation parameters algorithm settings will open towards the bottom of the menu panel. Manually generate the parameters to use for all replicates by clicking the blue arrow button at the bottom of the interface. Ensure the correct source channel is selected and check the &#39;Smooth&#39; box.
	<ol>
		<li>Set the surface detail to 0.7 &#181;m and select Background Subtraction (Local Contrast). Type the average cell length into the &#39;Diameter of the largest Sphere which fits into the Object&#39; box. Press the same blue arrow at the bottom when finished. 
		NOTE: This value can be estimated using the &#39;Slice&#39; tab in the toolbar and measuring the width of average cells.</li>
	</ol>
	</li>
	<li>For thresholding, determine the intensity histogram where only the brightest cells are segmented. Using the slicer, move up and down through the image stack to ensure it is as accurate as possible.
	<ol>
		<li>Select Enable for &#39;Split touching objects (Region Growing)&#39; and set the Seed Points Diameter to the same diameter as used previously. Ensure &#39;Intensity Based&#39; thresholding is selected and click the blue right arrow button.</li>
	</ol>
	</li>
	<li>The next two steps, Filter Seed Points and Filter Surfaces, can be adjusted by several measurements to ensure surfaces generated are accurate; however, baseline analysis requires no additional filtering. When no change is needed, click on the blue arrow button. 
	NOTE: The final step allows further classification of the surface depending on the desired output. After any edits are made, click the green arrow button to finish creating the surfaces.</li>
	<li>To save the Creation Parameters for batch analysis, click on the wand icon Creation. Click on Store Parameters for Batch and name it, then click on Ok. Representative processed images for t = 0 and t = 24 h are shown in Figures 2C,D, respectively. 
	NOTE: All objects within a selected &#39;Surface&#39; can be classified into sets based on various properties given by the image conversion software, such as surface area, volume, intensity, and distance from other created surfaces.</li>
	<li>Surface properties are found by selecting the Statistics tab. To gather all cell heights, click on the Detailed tab and select Specific Values and Position Z from the successive dropdown menus. Click on the single Save icon to save all z-positions and any classifications made into a .xls file. Repeat for all images.</li>
	<li>Find the median z-position of non-migrating cells from the representative image and subtract all z-positions below that number from each test condition well. 
	NOTE: Migration values are reported as the means of medians for each technical replicate condition above non-migrating cell height. They can be reported as median height, seen in Figure 3A, or as fold change in migration height for the desired timepoint compared to t = 0, seen in Figure 3B.</li>
</ol>
4. Parallel Layered Outward Spheroid Migration Assay (PLOSMA)method: Cell culture and hanging droplet culture for 3D spheroids
NOTE: This protocol describes cell culture and hanging drop culture adapted from the protocol authored by Nandi et al.14.
<ol>
	<li>Thaw HDFs according to the manufacturer&#39;s protocol. Passage as needed until the desired passage is achieved.</li>
	<li>In an aseptic cell culture hood, prepare a Petri dish by adding 10 mL PBS to the bottom and flipping over the lid so that the exterior rests atop the cell culture hood.</li>
	<li>Add the appropriate volume of cells (determined from cell count) to a microcentrifuge tube. Add cell tracking dye diluted in media. Bring the total volume to 1 mL with warmed media. 
	NOTE: Spheroids should be roughly 8000 cells per spheroid but may change based on cell type.</li>
	<li>Incubate the cell solution for 45 min at 37 &#176;C. Spin the cells down according to the manufacturer&#39;s recommended speed and aspirate the supernatant.</li>
	<li>Resuspend cells in 1:100 methylcellulose in media.</li>
	<li>Pipette 20 &#181;L droplets of the cell/media solution onto the lid of the Petri dish.</li>
	<li>Confidently, quickly, and carefully invert the lid and place it on top of the bottom half of the Petri dish containing PBS.</li>
	<li>Incubate the droplets for at least 24 h. 
	NOTE: Spheroid formation can be monitored by brightfield microscopy.</li>
</ol>
5. Parallel Layered Outward Spheroid Migration Assay (PLOSMA) method: Seeding cell spheroids onto granular hydrogel
NOTE: The following process is summarized in Figure 4A.
<ol>
	<li>To set up the PLOSMA method outlined in Figure 4A, aseptically add 15 &#181;L of gel using a positive displacement pipette to wells in a clear 96-well plate.</li>
	<li>Using a plate-spinning rotor centrifuge attachment, spin at 1000 x g for 10 s to flatten the gel. Flip the plate 180&#176; and spin again at 1000 x g for 10 s to ensure even gel distribution across the well bottom.</li>
	<li>Once even flatness has been achieved, aseptically photo-crosslink the gel from the top by applying focused light (365 nm, 33.4 mW/cm2) to the sample for 30 s to anneal the scaffold.</li>
	<li>Aseptically move the Petri dish of hanging droplets into the aseptic tissue culture hood and invert the lid.</li>
	<li>Using a 20 &#181;L pipette, slowly uptake a droplet until the spheroid enters the pipette tip. Eject droplet onto the scaffold in the center of the well.</li>
	<li>Repeat the previous steps for all wells. Ensure that each well has a spheroid by confirming with either brightfield or fluorescent microscopy.</li>
	<li>Incubate the well plate at 37 &#176;C for 2 h to allow spheroids to attach to the scaffold.</li>
	<li>Pipette an additional 15 &#181;L of gel on top of each spheroid. To ensure there is even gel distribution, centrifuge the plate at 300 x g for 15 s in each direction.</li>
	<li>Anneal the top layer of gel for 30 s using UV (365 nm) light at 33.4 mW/cm2. Pipette media on top of each scaffold to bring the total volume of the well to 200 &#181;L. 
	NOTE: Scaffolds will be very dry at this point, so add media dropwise down the side of the well to avoid detaching the spheroid.</li>
</ol>
6. Parallel Layered Outward Spheroid Migration Assay (PLOSMA) method: Confocal imaging of spheroids
NOTE: The ease of imaging depends on the imaging system. Locate the spheroid within the well at a low exposure time. The cells were imaged using the FITC channel (488 nm). The dye used for the cells had an excitation at 492 nm and an emission at 517 nm. 10x magnification provides increased detail over 4x magnification.
<ol>
	<li>Find the lowest stage level (z height) at which cells are still in focus. Set this as the lower limit of the z-stack.</li>
	<li>Find the highest stage level (z-height) at which cells are still in focus. Set this as the upper limit of the z-stack. 
	NOTE: Best imaging results include a step size of less than or equal to 5 &#181;m to maintain cell-scale resolution. There may be trade-offs between imaging speed and resolution depending on the confocal microscope system.</li>
	<li>Image all spheroids as described at t = 0 and 24 h. A 48-h timepoint may also be imaged depending on experimental constraints. Representative maximal projection images of t = 0 and t = 24 are seen in Figure 4B.</li>
</ol>
7. Parallel Layered Outward Spheroid Migration Assay (PLOSMA) method: 3D image analysis
<ol>
	<li>Import images into the analysis software as outlined in steps 3.1 through 3.4. In the top right corner of the left panel, click on the dropdown menu for Channel 1 and select Background Subtraction. Press Ok in the bottom of the panel.</li>
	<li>Once back in 3D view, press Auto Adjust all Channels in the Display Adjustment popup window and correct as needed.</li>
	<li>In the smaller toolbar just above the side menu, click on the Add new Reference Frame icon shown in Figure 5A with three orthogonal arrows to add a new tab called &#39;Reference Frame 1&#39;.</li>
	<li>Move the origin to the center of the spheroid in all three planes, as visualized in Figure 5B.</li>
	<li>In the same toolbar as the three orthogonal arrows, click on the icon with orange spheres and add new Spots to create a tab called &#39;Spots 1&#39;. Press the blue arrow button.</li>
	<li>Under Spot Detection, set the Estimated XY Diameter to the estimated diameter of the cells. Press the blue arrow button. 
	NOTE:&#160;For HDFs, this number is 15.0 &#181;m.</li>
	<li>For thresholding, adjust the intensity histogram to encircle only the brightest parts. Using the slicer, move up and down through the image stack to ensure it is as accurate as possible. Hit the blue next arrow.</li>
	<li>Press the green Execute button to finish the analysis.</li>
	<li>Uncheck Render on slicer or click on the yellow square icon on the right-hand side of the setup panel.</li>
	<li>Click on the Statistics tab. In the first dropdown menu, select Specific Values. In the second dropdown menu, select Distance from Origin Reference Frame. All values for the selected surfaces will be displayed, as seen in Figure 5C. Click the single save icon, which downloads a .xls file.</li>
	<li>Save the changes made to the image and the analyses by pressing the Save icon in the main toolbar. Figure 6A represents a 3D rendering of a spheroid imaged at 24 h, while Figure 6B represents the IMARIS Spots function marking spread cells, color-coded according to the distance from the origin reference frame.</li>
	<li>Normalize the exported data to the t = 0 images and calculate the mean of the cellular distance traveled and z-height for each spheroid to obtain a single value for each sample. Figure 7A,B depict representative graphs for each output, respectively.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+to+characterize+granular+hydrogel+rheological+properties,+porosity,+and+cell+invasion.">Methods to characterize granular hydrogel rheological properties, porosity, and cell invasion.</a> ACS Biomater Sci Eng. 8 (4), 1427-1442 (2022).</li><li>Shaik, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+matrix-derived+granular+hydrogel+enhances+cell+function+in+3D+culture.">Cardiac matrix-derived granular hydrogel enhances cell function in 3D culture.</a> ACS Appl Mater Interfaces. 16 (43), 58346-58356 (2024).</li><li>Puiggal&#237;-Jou, A., Asadikorayem, M., Maniura-Weber, K., Zenobi-Wong, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+factor-loaded+sulfated+microislands+in+granular+hydrogels+promote+hMSCs+migration+and+chondrogenic+differentiation.">Growth factor-loaded sulfated microislands in granular hydrogels promote hMSCs migration and chondrogenic differentiation.</a> Acta Biomater. 166, 69-84 (2023).</li></ol>

The authors have no conflicts of interest to disclose.

This protocol describes two in vitro models for characterizing cell migration in 3D for wound healing and tissue integration. The first model, the monolayer-based migration assay, relies on properly attached and confluent cells. This protocol was developed with a fibroblast cell type and optimized at a seeding density of 1,20,000 cells/cm2. This density allows cells to grow overnight to at least 80% confluency evenly across the bottom of the well plate. This step ensures migration in the z-direction within at least 24 h; if confluency is too low upon the addition of the gel layer, cells may continue to spread across the tissue culture plastic as well as into the gel, resulting in a non-uniform, slowed migration pattern, which was observed during optimization. Uneven migration heights may still be observed in areas of less dense cells, even at 80% confluency. Well replicates will reduce the noise of these cell behaviors. Overly confluent cells can cause the lifting of cells during the centrifugation period and potentially cell death. This variability is addressed by seeding at a consistent number of cells and by capturing a consistent image area to allow for appropriate data comparisons. To the author&#39;s knowledge, plate centrifugation has not been published for gel flattening, but centrifugation is commonly used for cell passaging and handling biomatter21,22. Adjusting the speed to match passaging speeds will maintain cell viability for further optimal cell processing.
The primary challenge in this method is maximizing imaging resolution and depth while minimizing imaging time to ensure the best analysis. Green cell tracking dye is sufficiently bright to image a 96-well with a 5 &#181;m or less step size and down to 1000 ms of exposure time. Lowering the exposure time reduces the amount of time cells are not in incubation conditions, but also reduces resolution. These parameters must be optimized on an individual microscope basis, but variability is reduced by ensuring all images are captured with the same settings within one study.
An important note for the analysis of MAMAs is that it requires eliminating the cells at or below the monolayer height to ensure only migrating cells are considered for statistical tests. Accordingly, the medians of replicate wells are reported due to the non-Gaussian distribution nature of the cell positions after filtering. Comparison between groups can be visualized with a histogram, and medians can be statistically analyzed with a non-parametric test.
Despite these challenges, the monolayer-based upward migration method is, at its simplest, a reproducible assay for 3D cell infiltration of porous scaffolds. To study the mechanistic effects of cell migration, ensure that parameters fit the cell type being studied. This may include the addition of chemotactic or haptotactic components, within the gel or in the media. Human dermal fibroblast complete media include migratory chemokines, but other cell types that use more specific cues require adaptation of the assay accordingly. This assay does lend itself to testing multiple types of variables; however, the scope of these is not covered in this protocol. The MAMA provides a physiologically relevant environment analogous to cell movement from bulk tissue into an injected porous hydrogel in vivo.
For the PLOSMA method, placement of the spheroids in the center of the scaffold is critical to successful imaging and meaningful cell migration in three dimensions. The exact seeding of the spheroid in the center of the gel is dependent on the user. To this end, steadying the pipette at the barrel with the user&#39;s non-dominant hand assists in centering, and the effectiveness of the seeding position can be confirmed using brightfield or fluorescent microscopy. An off-center spheroid can be remedied by a second attempt with a new spheroid, either on the same scaffold or on a new scaffold. For this reason, the authors recommend creating more spheroids than necessary and preparing more MAP gel than necessary.
The second layer centrifugation step ensures that the spheroid is (1) covered evenly by the gel and (2) able to spread evenly upward and downward into the gel, which is crucial to studying delivered cells. Centrifugation can also cause the spheroid to move from the center toward the edges of the well, and while this protocol limits this phenomenon by optimizing centrifugation steps and volume of the gel used for each layer (15 &#181;L) for even distribution, it does not completely eliminate its movement. The exact centrifugation speed and timing required to reduce spheroid movement may need to be adjusted according to the model of the centrifuge; however, the specification described in this protocol may be used as a benchmark for individual optimization. Another approach is to allow the spheroids 2 h of incubation time to attach to the scaffold before adding the second layer of gel. Spheroid movement is mitigated particularly well when both strategies are implemented. Finally, because of the multi-step centrifugation process, this method may not be suitable for less hardy cell lines.
Apart from the logistics of plating the spheroids in the PLOSMA method, there are limitations during image acquisition. The spheroid can be imaged using 4x or 10x magnification, but for best results, use at least a 10x magnification and reduce the step size of the z-stacks to 2-5 &#181;m. Magnification should be consistent throughout the study. Imaging time increases with higher resolution, so limit the number of samples in each well plate (4-8 wells per plate) to minimize time outside of the incubator. A live-imaging setup could also improve tracking and provide greater insights.
Because granular hydrogels have unique topology and design parameters that include inherent volume, porosity, mechanical strength, and, in some cases, bioactivity, it is necessary to study cell behavior in relation to these aspects with as much fidelity as possible. The PLOSMA method is designed to model cell movement after delivery or after cells have fully entered a granular gel. Because the cells are forced to migrate through the pores inherent in granular hydrogel geometry, the PLOSMA method effectively isolates porosity as an influence on cell behavior. Potential applications for this assay are cell delivery in situ and tissue integration within a granular scaffold, particularly in the wound healing space23.
Both protocols were developed with primary human dermal fibroblasts due to the role of fibroblast migration in tissue repair and remodeling4,24, however, the migratory behavior of any adherent cells may be measured in response to alteration of the porous scaffold - including additions of growth factors and surface/bulk composition of gel. These changes may require tailoring of these assays for appreciable results. Parameters requiring further optimization include cell seeding density, experiment duration, and/or analysis pipeline. IMARIS is a powerful imaging analysis tool that is utilized for cell migration analysis and has capabilities beyond what is outlined here, which include classifying all objects within a selected &#39;Surface&#39; into sets based on various properties such as surface area, volume, intensity, and distance from other created surfaces. There are many online resources to determine further analysis methods.
The two methods outlined here not only address the initial state of tissue introduction to a granular material in a physiological way, but also the subsequent cell response when fully embedded within the material. As with all migration assays, the cells present are capable of proliferating in parallel to movement, however the design of the described assays does not disrupt proliferation and thus ensures no undue impact on analysis. Both methods are compatible with endpoint staining in addition to longitudinal imaging, which uses PFA fixation to detect metrics such as cytoskeleton, collagen deposition, proliferation, and more. The use of the outlined methods moves towards a more accurate spatio-temporal representation of 3D cell migration that utilizes cell infiltration as a measurable parameter in contrast to previous methods1,6,14,15,25,26,27.

Biomaterials for therapeutic applications are increasingly evolving towards more complex and relevant models of cell environments to study tissue integration. Biomaterial scaffolds provide a three-dimensional (3D) structure for cell growth and aim to mimic a desired tissue1,2. Three-dimensional cell culture models include natural matrices and synthetic scaffolds that provide cells with further complexity via haptotactic or chemotactic cues3,4. Traditional hydrogel scaffolds are crosslinked in bulk, yielding a nanoporous mesh that allows diffusion of small molecules5,6, but requires degradation for cell-scale migration into a tissue area in need of repair7. Granular hydrogels are a subset of biomaterials that have a high potential for clinical translation due to their biocompatibility, ability to conform to irregular shapes, and, in many cases, their injectability8,9. Their building-block nature provides the advantage of cell-scale porosity to enhance tissue infiltration and angiogenesis as well as modularity, which allows for the addition of heterogeneous cues for cell behavior10,11,12. Understanding cell response and movement within a 3D scaffold is vital for physiological relevance in all applications using biomaterials for tissue integration.
Studying tissue ingrowth in three dimensions, however, has proven to be difficult to accomplish with quantitative accuracy. The expanded complexity of a 3D environment requires in vitro models of cell migration that can not only provide insight into cell behavior but also material condition optimization. Previously published reports of 3D granular scaffold cell migration have used topical seeding to explore cell behavior, reporting infiltration into the porous structure and cell morphology13 and others spheroid sprouting14,15, measuring outgrowth length and number of sprouts. Topical seeding migration lengths may be unevenly influenced by gravity forces, and due to microscopy limitations, results cannot be longitudinal. The spheroid sprouting method has been limited to 2D quantification via maximal projection, which is unable to capture the mechanism of controlled invasion. Both methods are measured in an xy-plane, which lacks the nuance necessary to fully recapitulate 3D cellular movement and scaffold infiltration.
This protocol describes two approaches for quantifying cell migration, such as in vitro infiltration into the porous 3D granular hydrogel scaffolds, specifically using Microporous Annealed Particle (MAP) scaffolds16,17,18,19. The purpose of the following methods is to study cell behavior in granular gels by controlling the directionality of their migration for three-dimensional analysis. The first, monolayer-based ascending migration assay (MAMA) approach, is a simplified model of endogenous cell integration that illustrates uniform cell-material interactions and serves as a platform to represent the initial environment in which cells interface with granular hydrogels as well as to isolate individual behavior prior to infiltration of the scaffold. The second, called the parallel layered outward spheroid migration assay (PLOSMA) method, is a 3D cell spheroid migration assay, that explores cell movement when fully surrounded by a complex scaffold environment and models cell movement after delivery, as well as movement after cells have fully entered a granular gel.
Both methods are quantifiable by 3D image analysis and can be applied to studying and optimizing material-cell interactions using longitudinal timepoints for regenerative medicine and tissue engineering applications, where promoting or restricting cell movement is within the design criteria. Additionally, these methods take advantage of plate centrifugation for uniform multi-well assay preparation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa Fluor 647 Phalloidin</td>
			<td>ThermoFisher</td>
			<td>A22287</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>VWR International</td>
			<td>332</td>
			<td></td>
		</tr>
		<tr>
			<td>CellTracker Green CMFDS Dye, 1 mg</td>
			<td>ThermoFisher</td>
			<td>C2925</td>
			<td>20 x 50 ug units, 492/517 nm</td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>ThermoFisher</td>
			<td>75016085</td>
			<td>ST Plus Series</td>
		</tr>
		<tr>
			<td>Clear 96 well plate</td>
			<td>MilliPore Sigma</td>
			<td>CLS3997-50EA</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Fisher Scientific</td>
			<td>MT-25950CQC</td>
			<td>250 mL</td>
		</tr>
		<tr>
			<td>Fibroblast Basal Medium</td>
			<td>ATCC</td>
			<td>PCS-201-030</td>
			<td>480 mL, phenol-red-free</td>
		</tr>
		<tr>
			<td>Fibroblast Growth Kit - Low Serum</td>
			<td>ATCC</td>
			<td>PCS-201-041</td>
			<td>7.5 mM L-glut,5 ng/mL rh FGF basic, 5 ug/mL rh Insulin, 1 ug/mL Hydrocortisone, 50 ug/mL Ascorbic acid, 2% FBS</td>
		</tr>
		<tr>
			<td>FIJI (ImageJ)</td>
			<td>NIH</td>
			<td></td>
			<td>Public access download</td>
		</tr>
		<tr>
			<td>Human Dermal Fibroblasts</td>
			<td>ATCC</td>
			<td>PCS-201-012</td>
			<td>Adult human dermal fibroblasts</td>
		</tr>
		<tr>
			<td>ImageXpress Micro Confocal</td>
			<td>Molecular Devices</td>
			<td></td>
			<td>Spinning Disc confocal microscope with 4x, 10x magnifications</td>
		</tr>
		<tr>
			<td>IMARIS</td>
			<td>Oxford Instruments</td>
			<td></td>
			<td>3/4D Imaged Visualizaiton and Analysis Software, Proprietary</td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>ThermoFisher</td>
			<td>Finnpipette F2 Variable volume Pipettes</td>
			<td>HeraCell Vios 160i CO2 Incubator, 165L</td>
		</tr>
		<tr>
			<td>M-20 Microplate Swinging Bucket Rotor</td>
			<td>ThermoFisher</td>
			<td>75003624</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylcellulose</td>
			<td>Fisher Scientific</td>
			<td>9004-67-5</td>
			<td>Lab grade, powder form</td>
		</tr>
		<tr>
			<td>Microcentrifuge tube</td>
			<td>Fisherbrand</td>
			<td>05-408-129</td>
			<td>1.5 mL microcentrifuge tubes</td>
		</tr>
		<tr>
			<td>Paraformaldehyde (4%)</td>
			<td>Alfa Aesar</td>
			<td>AAJ19943K2</td>
			<td>For fixing&#160;</td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Corning</td>
			<td>08-757-100A</td>
			<td>Bacteriological Petri Dishes with Lid 35 x 10 mm</td>
		</tr>
		<tr>
			<td>Pipettes</td>
			<td>ThermoFisher</td>
			<td>4642080</td>
			<td>Finnpipette F2 Variable volume Pipettes</td>
		</tr>
		<tr>
			<td>Sterile PBS</td>
			<td>Gibco</td>
			<td>10010-023</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton-X</td>
			<td>Fisher Scientific</td>
			<td>327371000</td>
			<td></td>
		</tr>
	</tbody>
</table>

quantifying three-dimensional cell migration within and into granular hydrogel biomaterials

Granular hydrogel scaffolds hold significant potential in regenerative medicine, functioning either as carriers for cell delivery or as interfaces for tissue integration. This article introduces two novel approaches for quantifying cell migration within and into granular hydrogels, highlighting the distinct applications of these scaffolds. First, a cell monolayer interface assay that simulates tissue growth into granular hydrogels for integration purposes is presented. Second, a spheroid-based assay is described, designed to track cell movement within the hydrogel matrix, specifically suited for applications involving cell delivery. Both methods enable precise and controlled measurements of cell migration, providing a comprehensive toolkit for researchers utilizing granular hydrogel scaffolds. The motivation for these methods stems from the need for tailored control over cell migration within the scaffold to align with specific applications. By optimizing and standardizing these quantification techniques, researchers can iteratively refine granular hydrogel properties, ensuring their effectiveness in diverse regenerative medicine contexts. This robust set of quantitative tools offers new opportunities to enhance granular hydrogel scaffolds, advancing their use in both cell delivery and tissue integration applications.

This protocol aims to detail the necessary steps for two novel granular scaffold migration assays. The MAMA method can be utilized to evaluate cellular infiltration at a tissue interface. Granular hydrogels are a more complex system than bulk hydrogels and, therefore, are inherently more complex to process for migration9,20. It is important to understand the stepwise process outlined in Figure 1. Each step builds on the next and has been optimized in this protocol. Seeding HDFs at a density of 120,000 cells/cm2 will result in at least 80% confluence overnight (Figure&#160;1A), and these non-fluorescent cells are best tagged with cell tracking dye the day of the experiment to maximize imaging potential (Figure&#160;1B). This protocol matches the lower centrifugal force used in HDF passaging to maintain cell viability. Due to the angle that is produced for a single centrifugation step, it is necessary to flip the plate 180&#176; to ensure gel shifts to fully cover the bottom surface of the wells (Figure&#160;1C). Allowing cells to recover in the incubator for 30 min after annealing (Figure&#160;1D) will maintain cell viability and result in optimal migration (Figure&#160;1E). A large area of a 96-well plate can be imaged with a 4x objective and matched from timepoint 0-24 h (Figure 2A,B) to widely assess cell behavior. Processing the resulting z-stack images in the analysis software provides advanced analyses for multiple large datasets in an easy-to-use interface. This protocol summarizes the steps to create datasets for cell height, or position Z, at each timepoint visualized with the representative images in Figure 2B,C. Analysis of the processed data is seen in Figure 3A, visualized using the mean of median heights and their standard deviation of each timepoint, and fold change height from non-migrating cells at t = 0 h is shown in Figure 3B. The underlying data of this method is typically non-normally distributed, so medians are more robust measures for comparison and are therefore used to summarize the data.
Likewise, the PLOSMA method can be utilized to evaluate the motility of delivered cells within a 3D granular hydrogel scaffold. Figure 4A outlines the steps to the PLOSMA method, and it is especially important to seed the spheroid in the center of the well. Centering the spheroid in the field of view is recommended but depends on the microscope&#39;s capabilities. Figure 4B displays representative images of spheroid spreading for t = 0 h and t = 24 h taken at 10x magnification in the FITC channel (488 nm). In the software, an origin reference frame can be created and adjusted to each z-stack (Figure 5A,B). The software can track radial distance from that origin reference frame and export it as the desired dataset (Figure 5C). Figure 6A shows a representative image of the 3D rendering of the t = 24 h spheroid, while Figure 6B shows the software&#39;s Spots function. An example of the processed data is shown in Figure 7. Figure 7A represents the average distance traveled from the center normalized to the Day 0 distances. Figure&#160;7B isolates distances traveled in just the z-plane since that is the direction the PLOSMA method aims to study.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67627/67627fig01.jpg" /> 
Figure 1:&#160;Monolayer-based Ascending Migration Assay cell culture and imaging. Schematic of major steps in cell and gel processing for MAMA. (A) Cells are grown to confluence overnight, and (B) cell tracking dye is added just prior to the addition of granular gel. The scaffold is assembled via (C) plate centrifugation and stabilized with (D) photo-crosslinking. Imaging at multiple timepoints allows (E) visualization of upward cell migration. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67627/67627fig02.jpg" /> 
Figure 2: MAMA image processing. Representative images of image processing. Top and side view comparison of raw confocal images at (A) t = 0 and (B) t = 24 h in the FITC channel on 4x magnification. (B) Top and side view comparison of the processed cell position Z heights at (C) t = 0 and (D) t = 24 h. Processing for 24 h includes subtraction of median non-migrating z-heights. Scale bars = 500 &#181;m. Abbreviations: MAMA = Monolayer-based Ascending Migration Assay. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67627/67627fig03v2.jpg" /> 
Figure 3: MAMA cell migration output analysis. (A) Median position Z and standard deviation of cell heights in each replicate (n = 6) at timepoints t = 0 (27.0 &#181;m &#177; 1.4 &#181;m) and t = 24 h (46.6 &#181;m &#177; 10.8 &#181;m). (B) Migration of cells at 24 h normalized to 0 h and reported as fold change (1.8 &#177; 0.4). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig03largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67627/67627fig04.jpg" /> 
Figure 4: Parallel Layer Outward Spheroid Migration Assay (PLOSMA) cell culture and imaging. (A) Schematic describing steps of scaffold layering. (B) Max intensity projections of spheroid taken at 0 and 24 h. Images were taken via confocal fluorescent microscopy in the FITC channel (488 nm) at 10x magnification. Scale bars = 200 &#181;m. Abbreviations: PLOSMA = Parallel Layer Outward Spheroid Migration Assay. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/67627/67627fig05.jpg" /> 
Figure 5: Creating a new origin reference frame. (A) The new origin reference frame button outlined with a red box. (B) The new origin is set to be in the center of the spheroid in all three dimensions. (C) Output metrics shown are the distances of cell surfaces from the origin reference frame, which describes how far cells have migrated from the center. Scale bar = 120 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/67627/67627fig06.jpg" /> 
Figure 6: 3D renderings of the embedded spheroid at 24 h. (A) Processed spheroid in 3D space. (B) The center of the same spheroid was determined using the origin reference frame function in IMARIS, and cell spread is color-coded by distance from the origin. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/67627/67627fig07.jpg" /> 
Figure 7: Example outputs for PLOSMA. (A) Example PLOSMA results showing distance traveled in &#181;m. The average distance traveled was 240.8 &#181;m &#177; 36.87 &#181;m. (B) Z-height fold change (tf/t0) of spheroid sprouting. The average fold change was 3.82 &#177; 1.495. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67627/67627fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

A protocol for the quantitative evaluation of 3D cell migration within and into the interface of granular hydrogels is presented here.

Quantifying Three-Dimensional Cell Migration Within and Into Granular Hydrogel Biomaterials

Granular hydrogel scaffolds hold significant potential in regenerative medicine, functioning either as carriers for cell delivery or as interfaces for ...

quantifying-three-dimensional-cell-migration-within-into-granular

Research

JoVE Journal

Bioengineering

386 Views.  University of Virginia. A protocol for the quantitative evaluation of 3D cell migration within and into the interface of granular hydrogels is presented here.

Video: Quantifying Three-Dimensional Cell Migration Within and Into Granular Hydrogel Biomaterials

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Alginate Hydrogels for Three-Dimensional Organ Culture of Ovaries and Oviducts

Ovarian cancer is the fifth leading cause of cancer deaths in women and has a 63% mortality rate in the United States1. The cell type of origin for ovarian cancers is still in question and might be either the ovarian surface epithelium (OSE) or the distal epithelium of the fallopian tube fimbriae2,3. Culturing the normal cells as a primary culture in vitro will enable scientists to model specific changes that might lead to ovarian cancer in the distinct epithelium, thereby definitively determining the cell type of origin. This will allow development of more accurate biomarkers, animal models with tissue-specific gene changes, and better prevention strategies targeted to this disease.
Maintaining normal cells in alginate hydrogels promotes short term in vitro culture of cells in their three-dimensional context and permits introduction of plasmid DNA, siRNA, and small molecules. By culturing organs in pieces that are derived from strategic cuts using a scalpel, several cultures from a single organ can be generated, increasing the number of experiments from a single animal. These cuts model aspects of ovulation leading to proliferation of the OSE, which is associated with ovarian cancer formation. Cell types such as the OSE that do not grow well on plastic surfaces can be cultured using this method and facilitate investigation into normal cellular processes or the earliest events in cancer formation4.
Alginate hydrogels can be used to support the growth of many types of tissues5. Alginate is a linear polysaccharide composed of repeating units of &beta;-D-mannuronic acid and &alpha;-L-guluronic acid that can be crosslinked with calcium ions, resulting in a gentle gelling action that does not damage tissues6,7. Like other three-dimensional cell culture matrices such as Matrigel, alginate provides mechanical support for tissues; however, proteins are not reactive with the alginate matrix, and therefore alginate functions as a synthetic extracellular matrix that does not initiate cell signaling5. The alginate hydrogel floats in standard cell culture medium and supports the architecture of the tissue growth in vitro.
A method is presented for the preparation, separation, and embedding of ovarian and oviductal organ pieces into alginate hydrogels, which can be maintained in culture for up to two weeks. The enzymatic release of cells for analysis of proteins and RNA samples from the organ culture is also described. Finally, the growth of primary cell types is possible without genetic immortalization from mice and permits investigators to use knockout and transgenic mice.

Culture of normal cells in their three-dimensional context represents an alternative method to study early events required for cellular transformation and tumorigenesis. This method is used to grow normal ovarian and oviductal cells to study early events in ovarian cancer formation.

Ovarian cancer is the fifth leading cause of cancer deaths in women and has a 63% mortality rate in the United States1. The cell type of origin for ...

Shape Memory Polymers for Active Cell Culture

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent shape upon the application of a stimulus such as heat1-5. In a typical shape memory cycle, the SMP is first deformed at an elevated temperature that is higher than its transition temperature, Ttrans [either the melting temperature (Tm) or the glass transition temperature (Tg)]. The deformation is elastic in nature and mainly leads to a reduction in conformational entropy of the constituent network chains (following the rubber elasticity theory). The deformed SMP is then cooled to a temperature below its Ttrans while maintaining the external strain or stress constant. During cooling, the material transitions to a more rigid state (semi-crystalline or glassy), which kinetically traps or "freezes" the material in this low-entropy state leading to macroscopic shape fixing. Shape recovery is triggered by continuously heating the material through Ttrans under a stress-free (unconstrained) condition. By allowing the network chains (with regained mobility) to relax to their thermodynamically favored, maximal-entropy state, the material changes from the temporary shape to the permanent shape.
Cells are capable of surveying the mechanical properties of their surrounding environment6. The mechanisms through which mechanical interactions between cells and their physical environment control cell behavior are areas of active research. Substrates of defined topography have emerged as powerful tools in the investigation of these mechanisms. Mesoscale, microscale, and nanoscale patterns of substrate topography have been shown to direct cell alignment, cell adhesion, and cell traction forces7-14. These findings have underscored the potential for substrate topography to control and assay the mechanical interactions between cells and their physical environment during cell culture, but the substrates used to date have generally been passive and could not be programmed to change significantly during culture. This physical stasis has limited the potential of topographic substrates to control cells in culture.
Here, active cell culture (ACC) SMP substrates are introduced that employ surface shape memory to provide programmed control of substrate topography and deformation. These substrates demonstrate the ability to transition from a temporary grooved topography to a second, nearly flat memorized topography. This change in topography can be used to control cell behavior under standard cell culture conditions.

A method for developing cell culture substrates with the ability to change topography during culture is described. The method makes use of smart materials known as shape memory polymers that have the ability to memorize a permanent shape. This concept is adaptable to a wide range of materials and applications.

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent ...

Planar and Three-Dimensional Printing of Conductive Inks

Printed electronics rely on low-cost, large-area fabrication routes to create flexible or multidimensional electronic, optoelectronic, and biomedical devices1-3. In this paper, we focus on one- (1D), two- (2D), and three-dimensional (3D) printing of conductive metallic inks in the form of flexible, stretchable, and spanning microelectrodes.
Direct-write assembly4,5 is a 1-to-3D printing technique that enables the fabrication of features ranging from simple lines to complex structures by the deposition of concentrated inks through fine nozzles (~0.1 - 250 &mu;m). This printing method consists of a computer-controlled 3-axis translation stage, an ink reservoir and nozzle, and 10x telescopic lens for visualization. Unlike inkjet printing, a droplet-based process, direct-write assembly involves the extrusion of ink filaments either in- or out-of-plane. The printed filaments typically conform to the nozzle size. Hence, microscale features (&lt; 1 &mu;m) can be patterned and assembled into larger arrays and multidimensional architectures.
In this paper, we first synthesize a highly concentrated silver nanoparticle ink for planar and 3D printing via direct-write assembly. Next, a standard protocol for printing microelectrodes in multidimensional motifs is demonstrated. Finally, applications of printed microelectrodes for electrically small antennas, solar cells, and light-emitting diodes are highlighted.

Planar and three-dimensional printing of conductive metallic inks is described. Our approach provides new avenues for fabricating printed electronic, optoelectronic, and biomedical devices in unusual layouts at the microscale.

Printed electronics rely on low-cost, large-area fabrication routes to create flexible or multidimensional electronic, optoelectronic, and biomedical ...

Three-dimensional Cell Culture Model for Measuring the Effects of Interstitial Fluid Flow on Tumor Cell Invasion

The growth and progression of most solid tumors depend on the initial transformation of the cancer cells and their response to stroma-associated signaling in the tumor microenvironment 1. Previously, research on the tumor microenvironment has focused primarily on tumor-stromal interactions 1-2. However, the tumor microenvironment also includes a variety of biophysical forces, whose effects remain poorly understood. These forces are biomechanical consequences of tumor growth that lead to changes in gene expression, cell division, differentiation and invasion3. Matrix density 4, stiffness 5-6, and structure 6-7, interstitial fluid pressure 8, and interstitial fluid flow 8 are all altered during cancer progression.
Interstitial fluid flow in particular is higher in tumors compared to normal tissues 8-10. The estimated interstitial fluid flow velocities were measured and found to be in the range of 0.1-3 &mu;m s-1, depending on tumor size and differentiation 9, 11. This is due to elevated interstitial fluid pressure caused by tumor-induced angiogenesis and increased vascular permeability 12. Interstitial fluid flow has been shown to increase invasion of cancer cells 13-14, vascular fibroblasts and smooth muscle cells 15. This invasion may be due to autologous chemotactic gradients created around cells in 3-D 16 or increased matrix metalloproteinase (MMP) expression 15, chemokine secretion and cell adhesion molecule expression 17. However, the mechanism by which cells sense fluid flow is not well understood. In addition to altering tumor cell behavior, interstitial fluid flow modulates the activity of other cells in the tumor microenvironment. It is associated with (a) driving differentiation of fibroblasts into tumor-promoting myofibroblasts 18, (b) transporting of antigens and other soluble factors to lymph nodes 19, and (c) modulating lymphatic endothelial cell morphogenesis 20.
The technique presented here imposes interstitial fluid flow on cells in vitro and quantifies its effects on invasion (Figure 1). This method has been published in multiple studies to measure the effects of fluid flow on stromal and cancer cell invasion 13-15, 17. By changing the matrix composition, cell type, and cell concentration, this method can be applied to other diseases and physiological systems to study the effects of interstitial flow on cellular processes such as invasion, differentiation, proliferation, and gene expression.

Interstitial fluid flow is elevated in solid tumors and can modulate tumor cell invasion. Here we describe a technique to apply interstitial fluid flow to cells embedded in a matrix and then measure its effects on cell invasion. This technique can be easily adapted to study other systems.

The growth and progression of most solid tumors depend on the initial transformation of the cancer cells and their response to stroma-associated signaling ...

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 ...

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. However in vivo, most of the cells are completely surrounded by the extracellular matrix (ECM), resulting in a three-dimensional (3D) distribution of receptors. This may trigger differences in the outside-in signaling pathways and thus in cell behavior.
This article shows that stimulating the dorsal receptors of cells already adhered to a 2D substrate by overlaying a film of a new material (a sandwich-like culture) triggers important changes with respect to standard 2D cultures. Furthermore, the simultaneous excitation of ventral and dorsal receptors shifts cell behavior closer to that found in 3D environments. Additionally, due to the nature of the system, a sandwich-like culture is a versatile tool that allows the study of different parameters in cell/material interactions, e.g., topography, stiffness and different protein coatings at both the ventral and dorsal sides. Finally, since sandwich-like cultures are based on 2D substrates, several analysis procedures already developed for standard 2D cultures can be used normally, overcoming more complex procedures needed for 3D systems.

The following protocol describes the procedure to assemble sandwich-like cultures to be used as an intermediate stage between bi-dimensional (2D) and three-dimensional (3D) cellular environments. The engineered systems can have applications in microscopy, biomechanics, biochemistry and cell biology assays.

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. ...

Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, which is regulated by a variety of soluble and physical signals in the microenvironment. Described here is a method created to study the process of branching morphogenesis by forming engineered three-dimensional (3D) epithelial tissues of defined shape and size that are completely embedded within an extracellular matrix (ECM). This method enables the formation of arrays of identical tissues and enables the control of a variety of environmental factors, including tissue geometry, spacing, and ECM composition. This method can also be combined with widely used techniques such as traction force microscopy (TFM) to gain more information about the interactions between cells and their surrounding ECM. The protocol can be used to investigate a variety of cell and tissue processes beyond branching morphogenesis, including cancer invasion.

This manuscript describes a soft lithography-based technique to engineer uniform arrays of three-dimensional (3D) epithelial tissues of defined geometry surrounded by extracellular matrix. This method is amenable to a wide variety of cell types and experimental contexts and allows for high-throughput screening of identical replicates.

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, ...

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

Currently, most in vitro models of wound healing, such as well-established scratch assays, involve studying cell migration and wound closure on two-dimensional surfaces. However, the physiological environment in which in vivo wound healing takes place is three-dimensional rather than two-dimensional. It is becoming increasingly clear that cell behavior differs greatly in two-dimensional vs. three-dimensional environments; therefore, there is a need for more physiologically relevant in vitro models for studying cell migration behaviors in wound closure. The method described herein allows for the study of cell migration in a three-dimensional model that better reflects physiological conditions than previously established two-dimensional scratch assays. The purpose of this model is to evaluate cell outgrowth via the examination of cell migration away from a spheroid body embedded within a fibrin matrix in the presence of pro- or anti-migratory factors. Using this method, cell outgrowth from the spheroid body in a three-dimensional matrix can be observed and is easily quantifiable over time via brightfield microscopy and analysis of spheroid body area. The effect of pro-migratory and/or inhibitory factors on cell migration can also be evaluated in this system. This method provides researchers with a simple method of analyzing cell migration in three-dimensional wound associated matrices in vitro, thus increasing the relevance of in vitro cell studies prior to the use of in vivo animal models.

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

The goal of this protocol is to evaluate the effect of pro- and anti-migratory factors on cell migration within a three-dimensional fibrin matrix.

Currently, most in vitro models of wound healing, such as well-established scratch assays, involve studying cell migration and wound closure on ...

Evaluation of Keratinocyte Proliferation on Two- and Three-dimensional Type I Collagen Substrates

Type I collagen, useful as a substrate for cell culture, exists in two forms: the two-dimensional, non-fibrous form and three-dimensional, fibril form. Both forms can be prepared with the same type I collagen. In general, the non-fibrous form promotes cell adhesion and proliferation. The fibril form (gels) provides more physiological conditions in many types of cells; therefore, gel culture is useful for examining physiological behaviors of cells, such as drug efficacy.
Researchers can select the appropriate form according to the purpose of its use. For example, in the case of keratinocytes, on-gel culture has been used as a wound healing model. FEPE1L-8, a keratinocyte cell line cultured on the non-fibrous form of type I collagen, promote cell adhesion. Notably, keratinocyte proliferation is slower on the fibril form than the non-fibrous form. Protocols for the preparation of type I collagen for cell culture are simple and have wide applications depending on the experimental needs.

Here, we present a protocol for the preparation of two different forms of culture substrates utilizing type I collagen. Depending on how collagen is handled, collagen molecules either maintain two-dimensional, non-fibrous form or reassemble into three-dimensional, fibril form. Cell proliferation on type I collagen is drastically affected by fibril formation.

Type I collagen, useful as a substrate for cell culture, exists in two forms: the two-dimensional, non-fibrous form and three-dimensional, fibril form. ...

Quantifying Three-Dimensional Cell Migration Within and Into Granular Hydrogel Biomaterials

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Chapters in this video

Three-dimensional Optical-resolution Photoacoustic Microscopy

Alginate Hydrogels for Three-Dimensional Organ Culture of Ovaries and Oviducts

Shape Memory Polymers for Active Cell Culture

Planar and Three-Dimensional Printing of Conductive Inks

Three-dimensional Cell Culture Model for Measuring the Effects of Interstitial Fluid Flow on Tumor Cell Invasion

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

Evaluation of Keratinocyte Proliferation on Two- and Three-dimensional Type I Collagen Substrates