A Customizable Chamber for Measuring Cell Migration

Summary

This protocol details a customizable method to measure cell migration in response to chemoattractants that may also be used to determine the diffusion rate of a drug out of a polymer matrix.

Abstract

Cell migration is a vital part of immune responses, growth, and wound healing. Cell migration is a complex process that involves interactions between cells, the extracellular matrix, and soluble and non-soluble chemical factors (e.g., chemoattractants). Standard methods for measuring the migration of cells, such as the Boyden chamber assay, work by counting cells on either side of a divider. These techniques are easy to use; however, they offer little geometric modification for different applications. In contrast, microfluidic devices can be used to observe cell migration with customizable concentration gradients of soluble factors^1,2. However, methods for making microfluidics based assays can be difficult to learn.

Here, we describe an easy method for creating cell culture chambers to measure cell migration in response to chemical concentration gradients. Our cell migration chamber method can create different linear concentration gradients in order to study cell migration for a variety of applications. This method is relatively easy to use and is typically performed by undergraduate students.

The microchannel chamber was created by placing an acrylic insert in the shape of the final microchannel chamber well into a Petri dish. After this, poly(dimethylsiloxane) (PDMS) was poured on top of the insert. The PDMS was allowed to harden and then the insert was removed. This allowed for the creation of wells in any desired shape or size. Cells may be subsequently added to the microchannel chamber, and soluble agents can be added to one of the wells by soaking an agarose block in the desired agent. The agarose block is added to one of the wells, and time-lapse images can be taken of the microchannel chamber in order to quantify cell migration. Variations to this method can be made for a given application, making this method highly customizable.

Introduction

In order for vital processes such as wound healing, immune responses, and embryonic development to occur, cell migration must take place. Cell migration involves the interaction between cells and neighboring cells, the extracellular matrix, and soluble chemical cues (attractants or repellants). As an example, in the process of wound healing, fibroblasts play an integral role in fibrogenesis and wound contraction, where the cells are recruited to the site of injury to synthesize collagen in order to form the extracellular matrix³. Numerous mechanisms behind the migration of fibroblasts to a wound site have been studied, and they include different mechanical, physical, electrical, and chemotactic factors⁴. Fibroblasts respond especially well to different concentration gradients of growth factors. These different growth factors work together to optimize tissue regeneration⁵. While observing the chemotactic response of fibroblasts to growth factor concentrations, one can study the pattern of directional migration of fibroblasts and how they orient themselves around physical obstacles in order to reach their destination. Therefore, the goals of this study were to first develop a system in which fibroblast growth could be tracked under guidance by physical barriers and to secondly model the growth of fibroblasts as they navigate through the system.

Currently, the Boyden chamber assay is the most widely used system to measure the migration of cells⁶. The Boyden chamber consists of a two-chamber multi-well plate where each well may contain medium with or without chemoattractants⁷. A filter membrane provides a porous interface between the two chambers in each well; this creates a barrier so that cells cannot pass through unless it is by active migration. Typically, for the Boyden chamber, a chemoattractant is added to the lower chamber, and the system is allowed to equilibrate to form a gradient between the upper and lower wells⁴. One problem with the Boyden chamber assay is that steep gradients end up forming along a single axis perpendicular with the surface of the membrane. This causes the difference in the chemoattractant concentration between the upper and lower wells to be a lower than what was originally expected. Due to this constraint, the Boyden chamber assay makes it hard to correlate specific cell responses with particular gradient characteristics, such as the slope and the concentration difference. Without these measurements, it is hard to study multi-gradient signal integration.

To address some of the constraints of the traditional Boyden chamber assay, microfluidic assays have been developed to form customizable concentration gradients¹. Standard methods for creating microfluidic systems require clean rooms for lithographic techniques. These techniques can be difficult to learn especially in a standard classroom setting. Thus, we have designed a chamber system for measuring cell migration that can be made without using a clean room. Using our system, the wells of the assay can be adjusted to a preferred size, and a linear concentration gradient of customized slope can be produced. This allows for accurate measurement of chemotaxis from random movement. The design is an inexpensive and easy-to-use system to model cell growth in response to different chemical stimuli.

Protocol

1. Producing a Microchannel Chamber to Create a Concentration Gradient

1. Cutting an acrylic mold piece
   1. Obtain a piece of acrylic of the desired width. Typically use 1/16-inch-thick acrylic sheets. Thicker sheets are difficult to cut well and very thin sheets do not have the requisite mechanical strength, causing them to break or warp during the process.
   2. Create a CAD file with the desired shape of the acrylic piece to produce a cavity within the polydimethylsiloxane (PDMS). Adjust the channel width and length to achieve the desired gradient relevant to individual experimental protocols.
      1. Here, use an acrylic piece of the following dimensions: two squares (0.254 cm x 0.254 cm) connected by a 0.127 cm wide channel (0.254 cm long) (Figure 1).
   3. Import the CAD file into the laser cutting apparatus and place the acrylic piece in a laser cutter according to the manufacturer's protocol.
      NOTE: Alternatively, 3D-print the piece from the CAD design. The advantage of this method is that alternate materials can be used for the mold; however, the resolution and reproducibility can be decreased with 3D printing compared to laser cutting.

Figure 1: Computer-Aided Design (CAD) Representation of Microchannel Chamber. This image depicts the CAD drawing of the acrylic insert needed to create the microchannel chamber. 1A) Top view of CAD drawing, 1B) Side view of CAD drawing with dimensions in inches, 1C) Top view of CAD drawing with dimensions in inches Please click here to view a larger version of this figure.

2. Pouring the polydimethylsiloxane (PDMS)
   1. In a weigh boat, mix 10 parts by weight of commercial Elastomer Base (e.g. Sylgard 184) with 1 part by weight of commercial Elastomer Curing Agent (e.g. Sylgard 194). Typically, mix 10 g of Elastomer Base to 1 g of Elastomer Curing Agent.
   2. Stir to combine the base and curing agent with a micropipette for 5 min.
   3. Set up a vacuum bell and place the weigh boat with PDMS in the vacuum for 30 min to remove trapped air bubbles.
   4. Turn off the vacuum and remove the weigh boat. Pour the PDMS on top of the acrylic cut-out in a small Petri dish. Make sure that the PDMS completely covers the insert.
   5. Allow the PDMS to cure overnight (at least 18 h) at room temperature.
3. Removing the insert from PDMS
   1. After curing the PDMS, remove the insert by carefully cutting the PDMS around the acrylic piece with a scalpel.

2. Plating the Cells in a Microchannel Chamber

1. Sterilizing the chamber for plating cells.
   NOTE: While PDMS can be sterilized using ethylene oxide or other techniques, a quick UV treatment is fast, inexpensive, and sufficient for cell culture studies. The short UV treatment duration did not impair the structure or mechanical integrity of the PDMS piece.
   1. In a standard cell culture cabinet, completely cover the chamber with 70% ethanol.
   2. Put the Petri dish in laminar flow hood with the lids off.
   3. Shut the hood and turn on the UV light. Expose to UV light for 1-2 h.
   4. Open the laminar flow hood and wait 15 min to reestablish flow.
   5. Remove the 70% ethanol.
   6. Wash the chambers twice with sterile phosphate buffered saline (PBS). Use 1 mL of PBS for each wash. Remove PBS and allow chambers to dry overnight with the lids off.
      NOTE: Alternatively, instead of using a laminar flow hood, use a UV chamber box for sterilization if one is available. Make a UV box by placing a UV light source in the back of a cardboard box lined with aluminum foil. The aluminum foil reflects the light to allow for even illumination of the sample.
2. Plating Cells in Chamber
   1. Count cells using Trypan blue and a hemocytometer.
      1. Add 100 µL of the cell suspension to 400 µL of 0.4% Trypan blue and mix gently. Apply 100 µL of this solution to the hemocytometer by gently filling both chambers under the glass coverslip.
      2. Use a microscope with a 10X objective to focus on the grid on the hemocytometer. Use a hand tally counter to count the live unstained cells within one set of 16 squares on the hemocytometer.
      3. Count cells in all 4 sets of 16 squares. Take the average cell count from the 4 sets of 16 squares and multiply by 5x10⁴. This final number is the number of cells/mL in the cell suspension.
   2. Add 2,500 cells to one well in each chamber. This translates to a near-confluent cell density of ~30,000/cm².
   3. Allow cells to sit in the chamber for 60 min before adding media (Dulbecco's Modified Eagle's Medium, 10% fetal bovine serum, 1% penicillin-streptomycin).

3. Dextran Soaked Agarose Blocks

1. Creating an agarose block
   1. Pour 3% agarose into the 3D printed mold (2 mm x 2 mm x 2 mm).
   2. Allow the agarose to solidify in a vacuum chamber for 30 min to remove any bubbles.
   3. Remove the agarose block from the mold.
      NOTE: Alternatively, pour 3% agarose into small Petri dish at a thickness of 2 mm. Cut a 2 mm x 2 mm x 2 mm block using a scalpel or other cutting tool. This is not as precise as the mold system but it can be a little easier to do.
2. Completely submerge the agarose block in a solution of the desired concentration of the chemotactic factor. To assess the concentration gradient formation, flush the chamber first with fluorescent dextran. The concentration and molecular weight of dextran should match that of a soluble factor or drug of interest.
3. Soak the agarose block overnight.

4. Time-lapse of the Dextran Diffusion to Assess the Soluble Factor Concentration Gradient

1. Place the dextran soaked agarose block in a small square well of the microchannel chamber.
2. Place the microchannel chamber in a cell imaging system or use another fluorescent microscope with a time-lapse capability. Begin the time-lapse according to manufacturer's instructions.
   NOTE: Results shown are taken with GFP filter, 50% light, 4/10 pH, and 4X magnification.

5. Time-lapse of Cell Growth

1. Plate cells in the microchannel chamber at one end of the chamber. Here, use 3T3 fibroblasts. Add 2,500 cells to one well in the microchannel chamber. Count cells as described in 2.2.1.
   1. Allow cells to sit in the chamber for 60 min before adding media (Dulbecco's Modified Eagle's Medium, 10% fetal bovine serum, 1% penicillin-streptomycin). Keep the microchannel chamber with plated cells at 37 °C in 5% CO₂. Cell migration through the channel can be viewed with a microscope.
   2. Add a marker or use a microscopic defect in the chamber as a marker in order to serve as a starting place to quantify the distance the cell front moves.
2. Establish a gradient by placing an agarose gel (8 mm³ block) containing the concentrated growth factor, chemo-attractant, drug, or other factor being tested (here, 40% FBS is being used as an example) at one end of the microchannel chamber.
3. Take time-lapse images of the moving cell front. Here, the results show images of the cell front that were taken every 12 h using an imaging system.
4. Use pictures of the cell front to quantify cell growth rate. Calculate the growth rate by first using the MATLAB polyfit function (roipoly) to create a polygonal mask defined by the cell front, the channel walls, and the reference marker. The dimensions of this mask yield the migration distance of the front from which average cell front velocity is calculated. Other studies have utilized a similar method⁸.
   NOTE: Compare the edge of the cell growth front on each frame to the concentration of the soluble factor at that same distance. The soluble factor concentration gradient is calculated using a 1D diffusion model approximation.
5. Take fluorescent images of the cells using Phalloidin and DAPI staining.
   1. Fix cells before staining with Phalloidin.
      1. Warm 4% paraformaldehyde at 37 °C.
      2. Add enough 4% paraformaldehyde to cover cells. Keep cells in the paraformaldehyde for 10 min.
      3. Rinse twice with PBS for 15 min each time. Use enough PBS to cover the cells.
   2. Remove PBS from cells and add enough permeabilizing solution (0.1% Triton X-100 in PBS) to cover cells. Leave solution for 10 min.
   3. Wash twice with PBS for 5 min each time. Use enough PBS to cover the cells.
   4. Remove PBS from cells and add blocking solution (1% bovine serum albumin in PBS) for 20 min.
   5. Remove blocking solution and wash 2 times with PBS for 5 min each time. Use enough PBS to cover the cells.
   6. From this point, protect cells from light with aluminum foil when not being used. Remove PBS from cells and add Phalloidin 488 diluted 1:50 in PBS for 1 h.
   7. Wash 2 times in PBS for 5 min for each wash. Use enough PBS to cover the cells. Then remove PBS from cells.
   8. Add DAPI (4',6-diamidino-2-phenylindole) diluted 1:1,000 in PBS to cells for 15 min.
   9. Remove DAPI and wash cells twice with PBS for 5 min for each wash.
   10. Remove last wash and add enough PBS to cover cells. Cells can now be imaged with a fluorescence microscope.

Results

Figure 2 shows the movement of the cell front across the channel in response to a gradient of fetal bovine serum placed at the opposite end of the channel from where the cells are plated. The cell front is shown at 48 h (Figure 2A), 72 h (Figure 2B), 96 h (Figure 2C), and 120 h (Figure 2D) post plating. The movement of the cell front was tracked with these time-lapse images, and the migration distance and...

Discussion

Our microchannel chamber may be used for a multitude of purposes, including determining the cell migration rate in response to growth factors and chemoattractants and measuring the diffusion rate of a drug from a polymer matrix. It is possible to utilize our microchannel chamber to grow cells and place a chemoattractant at one end of the chamber. The cells grow in response to the chemoattractant, and the cell migration can be quantified by taking time-lapse images that may be analyzed in order to determine the cell migra...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Sylgard 184 Silicone Elastomer Kit	Sigma-Aldrich	761036	poly(dimethylsiloxane) 2-part kit including silicone elastomer base and silicone elastomer curing agent
Acrylic Sheets	US Plastic	44200
Disposable Petri Dishes	Falcon	25373-041
Fluorescein isothiocyanate–dextran	Sigma-Aldrich	FD20s-100MG
Agarose, Type I, Low EEO	Sigma-Aldrich	A6013-100G
Dulbecco's Modified Eagle's Medium	Fisher Scientific	11965092	Cell media components
Fetal Bovine Serium	Fisher Scientific	16000036	Cell media components
Penicillin-streptomycin	Fisher Scientific	15140148	Cell media components
Phosphate Buffered Saline (PBS)	Fisher Scientific	BP24384
EVOS XL Cell Imaging System	Thermo Fisher Scientific	AME3300	Instrument used for taking time-lapse images
Versa Laser	Universal Laser Systems, Inc.	Model number VLS2.30	Laser cutter used for cutting plastic