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

Summary

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

Abstract

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.

Introduction

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 tissue^1,2. Three-dimensional cell culture models include natural matrices and synthetic scaffolds that provide cells with further complexity via haptotactic or chemotactic cues^3,4. Traditional hydrogel scaffolds are crosslinked in bulk, yielding a nanoporous mesh that allows diffusion of small molecules^5,6, 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 injectability^8,9. 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 behavior^10,11,12. 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 sprouting^14,15, 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) scaffolds^16,17,18,19. 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.

Protocol

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.

1. 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.
2. 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.
3. 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 µm sterile filter. Add this solution 1:1 v/v with the amount of gel necessary to conduct the experiment.
4. Incubate the 0.2 mM LAP, gel, and media solution at 37 °C on a tube rotator at 20 rpm for at least 30 min prior to proceeding to allow diffusion of LAP throughout the microparticles.
5. Once 30 min have elapsed, centrifuge the particle suspension at 18,000 x g for 5 min at 25 °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.

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.

1. Thaw human dermal fibroblasts (HDFs) according to the manufacturer's protocol. Passage as needed until the desired passage is achieved; generally, primary cells maintain genetic and phenotypic makeup through P5.
2. 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).
3. 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.
4. 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.
5. Add 20 µL 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.
6. Using a plate-spinning rotor centrifuge attachment at 25 °C, spin at 100 x g for 15 s with acceleration and deceleration of 8 to flatten the gel. Flip the plate 180° and spin again at 100 x g for 15 s to ensure even gel distribution across the well bottom, as seen in Figure 1C.
7. 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 µL of media to each well of cells after all scaffolds are formed. Let the cells incubate at 37 °C for 30 min to allow them to attach to the granular scaffold before imaging.
8. 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.
   1. Find the highest point of the cell fluorescent signal and set it as the upper edge of the z-stack. Use a 5 µm or smaller step size for best resolution.
9. 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.
10. 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.
11. 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.
12. 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 µL of sterile PBS twice for 5 min each. Add 200 µL of 4% Paraformaldehyde (PFA) for 20 min, then aspirate and discard. Wells can be stained immediately or stored at 4 °C in 1x PBS for up to a week.

3. Monolayer-based Ascending Migration Assay (MAMA) method: 3D image analysis

1. 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.
2. Open the software by double-clicking the desktop IMARIS Arena icon and selecting an image from the Arena.
3. The image is automatically loaded into the '3D View' analysis tab seen in the toolbar icon panel at the top. Click on the Image Proc tab in the main toolbar.
4. 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 '3D View'. Representative images for t = 0 h and t = 24 h are in Figure 2A,B, respectively.
5. 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 'Surfaces 1'.
6. 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 'Smooth' box.
   1. Set the surface detail to 0.7 µm and select Background Subtraction (Local Contrast). Type the average cell length into the 'Diameter of the largest Sphere which fits into the Object' box. Press the same blue arrow at the bottom when finished.
      NOTE: This value can be estimated using the 'Slice' tab in the toolbar and measuring the width of average cells.
7. 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.
   1. Select Enable for 'Split touching objects (Region Growing)' and set the Seed Points Diameter to the same diameter as used previously. Ensure 'Intensity Based' thresholding is selected and click the blue right arrow button.
8. 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.
9. 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 'Surface' 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.
10. 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.
11. 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.

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.

1. Thaw HDFs according to the manufacturer's protocol. Passage as needed until the desired passage is achieved.
2. 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.
3. 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.
4. Incubate the cell solution for 45 min at 37 °C. Spin the cells down according to the manufacturer's recommended speed and aspirate the supernatant.
5. Resuspend cells in 1:100 methylcellulose in media.
6. Pipette 20 µL droplets of the cell/media solution onto the lid of the Petri dish.
7. Confidently, quickly, and carefully invert the lid and place it on top of the bottom half of the Petri dish containing PBS.
8. Incubate the droplets for at least 24 h.
   NOTE: Spheroid formation can be monitored by brightfield microscopy.

5. Parallel Layered Outward Spheroid Migration Assay (PLOSMA) method: Seeding cell spheroids onto granular hydrogel

NOTE: The following process is summarized in Figure 4A.

1. To set up the PLOSMA method outlined in Figure 4A, aseptically add 15 µL of gel using a positive displacement pipette to wells in a clear 96-well plate.
2. Using a plate-spinning rotor centrifuge attachment, spin at 1000 x g for 10 s to flatten the gel. Flip the plate 180° and spin again at 1000 x g for 10 s to ensure even gel distribution across the well bottom.
3. 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.
4. Aseptically move the Petri dish of hanging droplets into the aseptic tissue culture hood and invert the lid.
5. Using a 20 µL pipette, slowly uptake a droplet until the spheroid enters the pipette tip. Eject droplet onto the scaffold in the center of the well.
6. Repeat the previous steps for all wells. Ensure that each well has a spheroid by confirming with either brightfield or fluorescent microscopy.
7. Incubate the well plate at 37 °C for 2 h to allow spheroids to attach to the scaffold.
8. Pipette an additional 15 µL 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.
9. 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 µL.
   NOTE: Scaffolds will be very dry at this point, so add media dropwise down the side of the well to avoid detaching the spheroid.

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.

1. Find the lowest stage level (z height) at which cells are still in focus. Set this as the lower limit of the z-stack.
2. 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 µm to maintain cell-scale resolution. There may be trade-offs between imaging speed and resolution depending on the confocal microscope system.
3. 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.

7. Parallel Layered Outward Spheroid Migration Assay (PLOSMA) method: 3D image analysis

1. 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.
2. Once back in 3D view, press Auto Adjust all Channels in the Display Adjustment popup window and correct as needed.
3. 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 'Reference Frame 1'.
4. Move the origin to the center of the spheroid in all three planes, as visualized in Figure 5B.
5. 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 'Spots 1'. Press the blue arrow button.
6. Under Spot Detection, set the Estimated XY Diameter to the estimated diameter of the cells. Press the blue arrow button.
   NOTE: For HDFs, this number is 15.0 µm.
7. 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.
8. Press the green Execute button to finish the analysis.
9. Uncheck Render on slicer or click on the yellow square icon on the right-hand side of the setup panel.
10. 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.
11. 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.
12. 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.

Discussion

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's knowledge, plate centrifugation has not been published for gel flattening, but centrifugation is commonly used for cell passaging and handling biomatter^21,22. 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 µm 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'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 µL) 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 µm. 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 remodeling^4,24, 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 'Surface' 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 methods^{1,6,14,15,25,26,27}.

Materials

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